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5,134 | https://bio-protocol.org/en/bpdetail?id=5134&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
A Bio-protocol resource
Peer-reviewed
An Autocatalytic Platform Combining a Nonlinear Hybridization Chain Reaction and DNAzyme to Detect microRNA
HZ Hongbo Zhang *
XC Xiuen Cao *
QZ Qubo Zhu
(*contributed equally to this work)
Published: Vol 14, Iss 24, Dec 20, 2024
DOI: 10.21769/BioProtoc.5134 Views: 293
Reviewed by: Han Deng Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in The Journal of Biological Chemistry Jun 2023
Abstract
MicroRNAs (miRNAs) are small, non-coding RNAs that play pivotal roles in gene regulation; they are increasingly recognized as vital biomarkers for various diseases, notably cancer. Conventional methods for miRNA detection, such as quantitative PCR and microarray analysis, often entail intricate sample preparation and lack the requisite sensitivity to detect low-abundance miRNAs like miRNA-21. This protocol presents an innovative approach that combines branched hybridization chain reaction (bHCR) with DNAzyme technology for the precise detection of miRNA-21. The bHCR amplifies the target signal through a branched structure, while the DNAzyme boosts detection sensitivity through catalytic cleavage, enabling swift and specific identification of miRNA-21. This dual amplification strategy offers a highly sensitive, specific, and rapid alternative to traditional techniques, making it particularly well-suited for early-stage disease diagnosis.
Key features
• This protocol enables a sensitive detection of miRNA-21.
• The technique employs an isothermal, enzyme-free bHCR process that can be carried out using standard laboratory equipment, eliminating the necessity for specialized instruments.
• The entire protocol can be finalized in less than five hours, offering a swift and effective approach for high-throughput miRNA detection.
• Minimal input RNA is needed for the protocol, and the sample preparation steps are straightforward.
Keywords: DNAzyme Branched hybridization chain reaction (bHCR) MicroRNA (miRNA) miRNA-21 Real-time quantitative polymerase chain reaction (RT-qPCR)
Graphical overview
Detection of microRNA-21 based on bHCR-DNAzyme platform.
Note: For a more detailed explanation of the construction and design principles of this platform, please refer to the previously published article [1].
Background
MicroRNAs (miRNAs) are a class of small non-coding RNAs that play a crucial role in gene expression regulation by targeting messenger RNA (mRNA) for degradation or translational repression, thereby influencing various cellular processes [2]. In numerous diseases, miRNA expression patterns are dysregulated. For instance, miR-21 is upregulated in various cancers, including liver, breast, lung, and colorectal cancer. Conversely, the let-7 family of miRNAs, recognized for their tumor-suppressive roles, is often downregulated in numerous cancer types [3]. As a result, miRNAs are significant biomarkers closely associated with the development of various diseases, including cancer [4–7]. Detecting nucleic acid biomarkers is crucial for disease identification, treatment, and intervention. However, traditional miRNA detection methods face challenges due to the low abundance and high sequence homology of nucleic acid biomarkers, as well as limited detection sensitivity and complex, time-consuming procedures [8,9]. These limitations significantly impede the speed of research and clinical diagnostics. Hence, there is an urgent need to develop a novel method that combines sensitivity, specificity, and operational simplicity.
This protocol introduces a novel detection strategy that combines the synergistic effects of branched hybridization chain reaction (bHCR) and DNAzyme technology. In bHCR, fuel probes quickly assemble into networked or branched nanostructures upon the presence of an initiator, leading to exponential signal amplification. This feature enables the sensitive detection of low-abundance molecules and accelerates reaction speed [10]. However, bHCR has limitations that restrict its application, such as an upper limit to amplification, signal saturation, and increased background signal due to the spontaneous opening and self-amplification of hairpin probes [11]. On the other hand, DNAzyme is a unique nucleic acid catalytic enzyme with biological catalytic activity, such as RNA (or DNA) cleavage and ligation [12], obtained through in vitro selection [13]. It is known for its ease of synthesis, modifiability, and cost-effectiveness. With its separation of the recognition sequence and catalytic core, DNAzyme allows for flexible sequence design. Moreover, DNAzyme-catalyzed nucleic acid amplification mimics enzymatic reactions, theoretically enabling unlimited signal amplification [14,15]. However, DNAzyme exhibits a relatively slow linear amplification rate, and its enzymatic reactions can be influenced by product accumulation and enzyme inactivation [12]. By combining bHCR and DNAzyme technologies, this protocol harnesses the strengths of both methods to overcome their individual limitations, resulting in a novel nucleic acid detection approach for miRNA. Table 1 provides a comparative analysis of the advantages and disadvantages of both techniques. Among various nucleic acid biomarkers, miRNA-21 is notably upregulated in several cancers. Its elevated levels are strongly associated with tumor progression, metastasis, and poor patient prognosis [16–18], highlighting its potential as both a biomarker and a therapeutic target for early cancer detection. Therefore, this protocol uses miRNA-21 as an example to demonstrate how the bHCR-DNAzyme platform can detect miRNAs from tissues or cells. This dual amplification system offers a highly sensitive and specific method for detecting miRNA-21, even at low concentrations, in complex biological samples. The integration of bHCR and DNAzyme addresses various constraints of current miRNA detection techniques, potentially leading to more reliable and user-friendly early cancer diagnostics. The outcomes of this study could have significant clinical implications, enhancing the accessibility of clinical diagnostics and disease monitoring.
Table 1. Overview of bHCR and DNAzyme
Branched hybridization chain reaction (bHCR) Deoxyribozyme (DNAzyme)
Concepts and principles By triggering DNA hairpin structures, branched double-stranded DNA can be formed, enabling exponential amplification of nucleic acids. A novel nucleic acid tool enzyme with biocatalytic activity, screened and isolated in vitro.
Advantages
• Isothermal amplification, requiring simple equipment.
• Enzyme-free amplification, low cost, easy reagent storage, good reproducibility.
• Exponential signal growth, rapid and strong signal output.
• Easy to synthesize, easy to modify, and cost-effective.
• Separate recognition sequence and catalytic center, offering flexible design.
• Comparable to enzymatic reactions, with no upper limit on signal amplification.
Disadvantages
• Amplification has an upper limit; signals can quickly saturate.
• Prone to spontaneous amplification, leading to high background signals.
• Linear reaction, slower speed.
• Product inhibition and enzyme inactivation require additional enzymes for long-term reactions.
Note: The principles of the two techniques are illustrated in Figure 1.
Figure 1. Principles of bHCR (A) and DNAzyme (B) techniques
Materials and reagents
Biological materials
Tumor and paratumor (PT) tissue from patients with hepatocellular carcinoma (from Third Xiangya Hospital in Central South University, Approval No: 2023009), stored in liquid nitrogen until analysis
Reagents
CaCl2 (Kermel Chemical Reagent Co Ltd., CAS: 10043-52-4)
NaCl (Kermel Chemical Reagent Co Ltd., CAS: 7647-14-5)
DEPC water (Biosharp, catalog number: BL510B)
HCl (Sinopharm, CAS: 7647-01-0)
1 M HEPES solution (Servicebio, catalog number: G4210-100ML)
Trizol (Jiangsu Cowin Biotech, catalog number: CW0580S)
Chloroform (Sinopharm, CAS: 67-66-3)
Isopropanol (Sinopharm, CAS: 67-63-0)
Ethanol (Sinopharm, Brand, CAS: 64-17-5)
MiRNA 1st strand cDNA synthesis kit (by tailing A) (Vazyme, Specification: 20rxns, catalog number: MR201-01)
Hieff® qPCR SYBR Green Master Mix (No Rox) (Yeasen, catalog number: 11201ES03)
10× TBE solution (Solarbio, catalog number: T1051)
30% Acr-Bis (29:1) (Coolaber, catalog number: SL1090)
10% APS (Coolaber, catalog number: SL1130)
TEMED (Coolaber, CAS: 110-18-9)
Labgreen (Coolaber, catalog number: SL2150)
6× loading buffer (Coolaber, catalog number: SL2210)
RNase-free water (Shanghai Promega, catalog number: SP119A)
Oligonucleotides (Beijing Tsingke Biotech, purification method: HPLC) (see Table 2 for details)
Table 2. Nucleic acid sequences used in this work
Name Sequence (5′-3′)
Trigger AGTCTAGGATTCAGCGTGGGATTA
S CTCGGCACAAGTGGGTACATT rA GAGTCTAGGATTCAGCGTGGGATTA
S1 CTCGGCACAAGTGGGTACATTAGAGTCTAGGATTCAGCGTGGGATTA
L TAATCCGACCCTGATATACCACTTGTGCCGAG
DNAzyme TAGACTTTCTCACAGCGTACTCGCTAAGGTTGTATGTAC
H1 TAATCCCACGCTGAATCCTAGACTCAAAGTAGTCTAGGATTCAGCGT GCGCTAAGGTTGTATGTAC
H2 AGTCTAGGATTCAGT(BHQ-1)CTAGGATTCAGCGTGCATCTCCACGCT GAATCCTAGACT(FAM)ACTTTG
H3 TAGACTTTCTCACAGCGGAGATGCACGCTGAATCCTAGACTTCCA GGAGTCTAGGATTCAGCGTG
H4 AGTCTAGGATTCAGCGTGGGATTACACGCTGAATCCTAGA CTCCTGGAAGCGTGGGATTA
miR-21 UAGCUUAUCAGACUGAUGUUGA
1-mut UAGCUUAUCAAACUGAUGUUGA
2-mut UAACUUAUCAGACUAAUGUUGA
4-mut UAGAUUAUCAAACUGAUAUUAA
H-miR-21 CTGAATCCTAGACTTCAACATCAGTCTGATAAGCTAAAAAGTCT AGGATTCAGCGTGGGATTA
Note: The fluorescent group FAM and the quencher group BHQ-1 are both modified on the thymine (T) base; rA indicates that the DNAs were replaced by RNAs. The underlined bases in 1-mut, 2-mut, and 4-mut are mutant bases.
Solutions
10 mM HEPES buffer (see Recipes)
75% ethanol solution (see Recipes)
1× TBE electrophoresis buffer (see Recipes)
2 µM SL initial concentration solution (see Recipes)
Recipes
10 mM HEPES buffer (100mL)
Reagent Final concentration Quantity
CaCl2 100 mM 1.110 g
NaCl 150 mM 0.877 g
1 M HEPES solution 10 mM n/a
DEPC H2O n/a n/a
Adjust the pH to 7.2 with HCl n/a n/a
Total (optional) n/a 100 mL
75% ethanol solution (10 mL)
Reagent Final concentration Volume
Anhydrous ethanol 75% (v/v) 7.5 mL
H2O n/a see note
Total n/a 10 mL
Note: The water is made using an ultrapure water machine.
1× TBE electrophoresis buffer (500 mL)
Reagent Final concentration Volume
10× TBE n/a 50 mL
H2O n/a see note
Total n/a 500 mL
Note: The water is made using an ultrapure water machine.
2 µM SL initial concentration solution (500 µL)
Reagent Final concentration Volume
100 µM S stock solutions 2 µM 2 µL
100 µM L stock solutions 2.5 µM 2.5 µL
DEPC H2O n/a 95.5 µL
Total n/a 100 µL
Note: Prepare 100 µM stock solutions of S and L according to the provided oligonucleotide synthesis instructions. For sequence information, manufacturer and catalog number, please refer to item 19 in the Reagents section. The preparation process is outlined in step B2 of the Procedure.
Laboratory supplies
0.2 mL microcentrifuge tube (Biosharp, catalog number: BS-02-P)
1.5 mL microcentrifuge tube (Biosharp, catalog number: BS-15-M)
2 mL microcentrifuge tube (Biosharp, catalog number: BS-20-M)
15 mL centrifuge tube (Servicebio, catalog number: EP-5001-J)
50 mL centrifuge tube (Servicebio, catalog number: EP-5001-J)
10 μL pipette tip (Servicebio, catalog number: P-10)
200 μL pipette tip (Servicebio, catalog number: TP-200)
1,000 μL pipette tip (Servicebio, catalog number: TP-1250)
Pipette (China Dalong Xingchuang Experimental Instrument Co., Ltd, model: TopPette)
Cell freezing tube, 2 mL (Corning, catalog number: CLS430659)
Equipment
High-speed freezing centrifuge (Changsha Yingtai, model: TGL16)
Ultrapure water machine (Bertone, model: K.L.MINI4-T)
PCR amplifier (Bio-Rad, model: Biometra)
Ultramicro ultraviolet spectrophotometer (Qua Well, model: Q5000)
Vortex mixer (Shanghai Huxi, model: XW-80A)
Quartz fluorescent micro cuvette, 1 mm (Wuxi edge spectrum optics)
Biochemical incubator (Shanghai Boxun medical, model: SPX-150B-Z)
Electrophoresis analyzer (Beijing Liuyi Instrument Factory, model: DYY-7C)
Vertical electrophoresis tank (Beijing Junyi Oriental electrophoresis equipment Co., Ltd, model: DYCP-31BN)
Electronic balance (Shanghai Guangzheng medical, model: YP-B2002)
Fluorescence spectrophotometer (HITACHI, model: F-7100)
Real-time fluorescent quantitative gene amplification instrument (Bio-Rad, model: CFX CONNECT)
Clean bench (Suzhou Zhijing purification equipment Co., Ltd, model: SW-CJ-2FD)
Chemiluminescence imager (Bio-Rad, model: ChemiDoc XRS+)
Micro analytical balance (Shimadzu, model: AUW120D)
Software and datasets
GraphPad Prism (version 7, GraphPad Software)
Procedure
Design the probe H0 of bHCR-DNAzyme using the NUPACK website
Note: The prediction of hairpin structures and interactions between hairpins was performed using the NUPACK website (http://www.nupack.org/).
The sequences of the probes Trigger, H1, H2, H3, H4, S, and L are fixed and do not require additional design. However, the H0 sequence must be designed each time based on the specific miRNA target being detected. For example, to design H0 for detecting miRNA-21 (referred to as H-miR-21), we constructed a DNA hairpin with a stem-loop structure. In this structure, bases 15–36 (in the 5' to 3' direction) form the loop and are fully complementary to the miRNA-21 sequence, while bases 40–63 correspond to the Trigger sequence.
Use the NUPACK website to validate the rationale behind the H0 design. Input the sequences of H-miR-21 and miRNA-21 into the website to predict their binding. If bases 15–36 (in the 5' to 3' direction) of H-miR-21 form complementary pairs with miRNA-21, thereby opening the hairpin stem and exposing the Trigger sequence, then the design of H-miR-21 can be considered successful.
Note: Based on the design strategy for H-miR-21, H0 can be modified for different miRNA sequences, allowing for the detection of other miRNAs or viruses. Previous studies have demonstrated that this platform can also detect mimics of COVID-19 and classical swine fever virus (CSFV).
Construction of the bHCR-DNAzyme platform
Prepare 10 mL of 10 mM HEPES buffer according to Recipes and store at 4 °C.
Note: All reactions and dissolution are carried out in this 10 mM buffer solution. The water used here is DEPC water, which is used for subsequent DNA and RNA oligonucleotide precipitation dissolution.
Prepare the bHCR-DNAzyme hairpin probe stock solutions: Dissolve all probe oligonucleotides in the buffer according to the synthesis instructions to prepare 100 µM stock solutions. Aliquot the stock solutions into 200 µL microcentrifuge tubes and store at -20 °C to avoid repeated freeze-thaw cycles that could degrade the DNA strands.
Note: All probe oligonucleotides, including Trigger, S, S1, H1, H2, H3, H4, miRNA-21, 1-mut, 2-mut, 4-mut, and H-miR-21, are listed in the nucleic acid sequences table under the Reagents section. The H2 hairpin probe is fluorescently labeled, so protect it from light during dissolution, storage, and reactions.
Prepare the 2 µM SL initial concentration solution: Mix the S and L stock solutions in a 4:5 ratio. Incubate the mixture at 25 °C in a PCR machine for 30 min to form the SL DNA duplex before use, achieving a final S concentration of 2 µM.
bHCR-DNAzyme nano self-assembly: Add different concentrations of Trigger solution to the 0.1 µM H1/H2/H3/H4/SL solution and incubate at 37 °C for 1 h.
Note: The different concentrations of Trigger mentioned here can be chosen by researchers according to their specific needs, or they can refer to section C. This section provides a straightforward description of the experimental steps involved in the platform's self-assembly process.
Validation of bHCR-DNAzyme Nano Self-Assembly
Non-denaturing polyacrylamide gel electrophoresis (PAGE) validation
Prepare the following solutions: 0.1 μM Trigger, 0.1 μM H1, 0.1 μM H2, 0.1 μM H3, 0.1 μM H4, 0.1 μM SL, 0.1 μM H1/H2/H3/H4, 0.1 μM H1/H2/H3/H4/SL, 0.1 μM Trigger/H1/H2/H3/H4/SL, 0.1 μM DNAzyme, 0.1 μM DNAzyme/SL, and 0.1 μM DNAzyme/S1L. Incubate at 37 °C for 1 h.
Note: S1 is the same as the S sequence but without RNA modification. The contents of step C1a are presented in Table 3.
Table 3. PAGE experiment solution groups
Group Component Group Component
1 0.1 μM Trigger 7 0.1 μM H1/H2/H3/H4
2 0.1 μM H1 8 0.1 μM H1/H2/H3/H4/SL
3 0.1 μM H2 9 0.1 μM Trigger/H1/H2/H3/H4/SL
4 0.1 μM H3 10 0.1 μM DNAzyme
5 0.1 μM H4 11 0.1 μM DNAzyme/SL
6 0.1 μM SL 12 0.1 μM DNAzyme/S1L
Prepare a 15% non-denaturing polyacrylamide gel: In a beaker, sequentially add 5 mL of 30% Acr-Bis (29:1), 2.5 mL of ultrapure water, 2.5 mL of 4× separating gel buffer, 100 μL of 10% APS, and 4 μL of TEMED, and mix well. Pour the solution into the gel casting plate, insert the gel comb, and let it solidify at room temperature for 20 min. Then, place the gel in an electrophoresis tank and add 400 mL of 1× TBE (see Recipes).
Note: The 4× separating gel buffer is obtained by diluting 10× TBE with ultrapure water.
Remove the gel comb and mix 10 μL of each solution prepared in C1a with 2 μL of 6× loading buffer; then, load the mixture into the wells.
Perform electrophoresis at a constant current of 20 mA for 90 min.
After electrophoresis, stain the gel with Labgreen for 30 min, then use a chemiluminescence imaging system for gel imaging to observe and record the experimental results.
Expected results: We anticipate that the band positions of group 7 will correspond to those of groups 2, 3, 4, and 5, indicating that the hairpins exist independently and have not been opened. Additionally, groups 9 and 11 are expected to produce bands with higher molecular weights and lower electrophoretic mobility compared to groups 8 and 12, confirming the success of self-assembly and the effective cleavage of SL by DNAzyme.
Fluorescence spectroscopy validation
Prepare the following solutions: 0.1 μM H2, 0.1 μM H1/H2/H3/H4, 0.1 μM H1/H2/H3/H4/SL, 50 nM Trigger + 0.1 μM H1/H2/H3/H4, and 1 nM/10 nM/50 nM Trigger + 0.1 μM H1/H2/H3/H4/SL. Incubate at 37 °C for 1 h.
Add each of the above solutions to a 1 mm quartz fluorescence microcuvette and measure the fluorescence intensity using a fluorescence spectrophotometer. Set the parameters as follows: slit width 5 nm, voltage 700 V, excitation wavelength 492 nm, emission wavelength range 500–600 nm.
Expected results: We anticipate a significant increase in the fluorescence intensity of the solution following the introduction of Trigger, with the intensity continuing to rise as the Trigger concentration increases. This will confirm the successful self-assembly of bHCR.
Detection performance of bHCR-DNAzyme for miR-21
Linear range and detection limit
H0 is designed to detect miRNA-21 and is referred to as H-miR-21. Dissolve the lyophilized H-miR-21 oligonucleotide in buffer according to the synthesis instructions to prepare a 100 μM stock solution, then dilute it to a 2 μM H-miR-21 working solution. Heat at 95 °C for 5 min and slowly cool to room temperature to form a hairpin structure. Store in the dark at 4 °C.
Prepare the following reaction systems separately: 0.01 nM miR-21 + 200 nM H1/H2/H3/H4 + 20 nM SL + 10 nM H-miR-21, 0.02 nM miR-21 + 200 nM H1/H2/H3/H4 + 20 nM SL + 10 nM H-miR-21, 0.05 nM miR-21 + 200 nM H1/H2/H3/H4 + 20 nM SL + 10 nM H-miR-21, 0.08 nM miR-21 + 200 nM H1/H2/H3/H4 + 20 nM SL + 10 nM H-miR-21, 0.1 nM miR-21 + 200 nM H1/H2/H3/H4 + 20 nM SL + 10 nM H-miR-21, and 0.2 nM miR-21 + 200 nM H1/H2/H3/H4 + 20 nM SL + 10 nM H-miR-21. Incubate at 30 °C for 1 h and measure the fluorescence intensity using a fluorescence spectrophotometer. Perform parallel detection on three samples for each group and include a blank control without miR-21. Set the parameters as follows: slit width 5 nm, voltage 700 V, excitation wavelength 492 nm, emission wavelength range 500–600 nm.
Specificity
Prepare a stock solution of miR-21 mutant oligonucleotide: Dilute the 100 μM miR-21 mutant stock solution with buffer to a final concentration of 5 μM and store at 4 °C.
Prepare the following reaction systems separately: 2 nM miR-21 + 200 nM H1/H2/H3/H4 + 20 nM SL + 10 nM H-miR-21, 2 nM 1-Mut + 200 nM H1/H2/H3/H4 + 20 nM SL + 10 nM H-miR-21, 2 nM 2-Mut + 200 nM H1/H2/H3/H4 + 20 nM SL + 10 nM H-miR-21, and 2 nM 4-Mut + 200 nM H1/H2/H3/H4 + 20 nM SL + 10 nM H-miR-21. Incubate at 30 °C for 1 h. Use the detection results from the 2 nM miR-21 group as the control and measure the fluorescence intensity using a fluorescence spectrophotometer. Perform parallel detection on three samples for each group and include a blank control without miR-21.
Total RNA extraction from tissues or cells
Note: The procedure for RNA extraction from tissues or cells has been clearly described in previous studies [19].
Treat all experimental equipment with solid-phase RNase inhibitor. Weigh 30 mg of tumor tissue (or 1.5 × 107–3 × 107 cells) and 30 mg of PT tissue, and place each in a mortar. Add 1 mL of Trizol and grind thoroughly. Let the mixture sit at room temperature for 5 min, then transfer it to a centrifuge tube and centrifuge at 1,609× g for 5 min.
Collect the supernatant and add 0.2 mL of chloroform. Shake for 15 s, let it sit at room temperature for 2 min, then centrifuge at 9,270× g for 15 min at 4 °C.
Caution: Use Trizol and chloroform in a well-ventilated area while wearing appropriate protective equipment, as both are hazardous chemicals.
Transfer 600 μL of the aqueous phase to a new centrifuge tube. Add 600 μL of isopropanol and mix thoroughly. Let the mixture sit at room temperature for 10 min, then centrifuge at 9,270× g for 10 min at 4 °C.
Discard the supernatant, wash the pellet with 1 mL of 75% ethanol, and centrifuge at 9,270× g for 3 min at 4 °C.
Note: Prepare the 75% ethanol with DEPC-treated water and pre-cool to 4 °C.
Discard the supernatant, air-dry the pellet at room temperature, and dissolve it in 20 μL of RNase-free water to obtain the total RNA solution.
Measure the total RNA concentration using a microvolume UV spectrophotometer and dilute it to 500 ng/μL.
Detection of miR-21 in tumor samples using bHCR-DNAzyme platform and RT-qPCR
Detection of miR-21 using bHCR-DNAzyme platform
Use the total RNA solution extracted in the previous step as the buffer to evaluate the detection capability of the bHCR-DNAzyme platform in biological samples. The experimental procedure follows the method described in section D.
Using the total RNA solution extracted in the previous step as the buffer, perform fluorescence detection of total RNA solutions from tumor tissue and PT tissue samples under the optimized conditions of the bHCR-DNAzyme platform. Perform parallel detection on three samples for each group and include a blank control.
Detection of miR-21 Using RT-qPCR
Reverse transcription: Use an RNA reverse transcription kit (miRNA 1st Strand cDNA Synthesis Kit). Mix 2 μg of total RNA (500 ng/μL, 4 μL), 10 μL of 2× miRNA RT Mix, 2 μL of HiScript miRNA Enzyme Mix, and 4 μL of RNase-free ddH2O to create a 20 μL reaction system. Mix well, then place it in a PCR machine and run the following program to obtain cDNA: 37 °C for 60 min, 85 °C for 5 min, and 4 °C for 5 min (see Table 4). Measure the concentration of cDNA using a microvolume UV spectrophotometer and dilute it to 50 ng/μL. Store at 4 °C.
Table 4. Thermocycling conditions for the reverse transcription
Step Temperature (°C) Duration Number of cycles
Step 1 37 60 min 1
Step 2 85 5 min 1
Step 3 4 5 min 1
Hold 4 ∞
RT-qPCR: Prepare a 20 μL reaction system with the following components: 10 μL of qPCR Mix, 7.2 μL of nuclease-free water, 2 μL of cDNA, 0.4 μL of forward primer, and 0.4 μL of reverse primer. Set up three parallel replicates for each sample. Mix thoroughly and perform the reaction in a real-time quantitative PCR machine under the following conditions: pre-denaturation at 95 °C for 5 min, denaturation at 95 °C for 10 s, annealing/extension at 60 °C for 30 s, for a total of 40 cycles (see Table 5). Finally, calculate and analyze the resulting Ct values using the 2-ΔΔCt method.
Note: Forward primer: 5'-ACACTCCAGCTGGGTAGCTTATCAGACTGA-3'; reverse primer: 5'-TGGTGTCGTGGAGTCG-3'.
Table 5. Thermocycling conditions for the PCR reaction
Step Temperature (°C) Duration Number of cycles
Pre-denaturation 95 300 s 1
Denaturation 95 10 s 40
Annealing/Extension 60 30 s
Hold 4 ∞ -
Data analysis
Linear range and selectivity of the bHCR-DNAzyme platform
The fluorescence intensity data of the sample was measured using a fluorescence spectrophotometer (F-7100) at 492 nm. The y-axis value is calculated by subtracting the fluorescence value of the blank control group from the average of the fluorescence values of three parallel samples. Figure 2A displays the variances in fluorescence intensity caused by miRNA-21 and various mutants. The results showed that the fluorescence intensities of the strands containing 1 (1-mut), 2 (2-mut), and 4 (4-mut) mutated bases were 34%, 27%, and 9%, respectively, of the fluorescence intensity of the miR-21 strand, demonstrating the platform's excellent specificity. Figure 2B presents the standard curve of miRNA-21 concentration versus fluorescence signal difference. The results also indicated a linear relationship between fluorescence intensity and miR-21 concentration in the range of 0.01 to 0.2 nM. The corresponding fitted equation is y = 695.52x + 4.112 (n = 3, R2 = 0.9944), with a detection limit of 0.003 nM. Statistical significance was evaluated using one-way ANOVA followed by Tukey’s post-hoc test. Data points were selected based on the results of the statistical analysis. Outliers were excluded if their deviation from the group mean exceeded three standard deviations or if they clearly fell outside the confidence interval of the fitted line.
Figure 2. bHCR-DNAzyme platform for (A) specificity and (B) sensitivity detection of miRNA-21. Error bar: SD, n = 3.
Comparison of the bHCR DNAzyme platform with the traditional RT-qPCR method
Figure 3 illustrates the detection outcomes of miRNA-21 in patients’ tumor tissues utilizing the bHCR-DNAzyme platform (Figure 3A) and the RT-qPCR platform (Figure 3B). It compares the expression levels of miRNA-21 between tumor and paratumor tissues. Statistical significance was assessed using one-way ANOVA followed by Tukey’s post-hoc test. The comparison between the two platforms underscores the enhanced accuracy of the bHCR-DNAzyme platform in discriminating tumor tissues from paratumor tissues, along with its superior error margin in contrast to the RT-qPCR platform. All patient samples were considered, with data points stemming from experimental irregularities or technical glitches being omitted.
Figure 3. Comparison of miR-21 detection results in patients' tissues detected by (A) bHCR-DNAzyme and (B) RT-qPCR. Error bar: SD, n = 3.
Note: Each experiment mentioned above involved three independent biological replicates and three technical replicates. This setup ensures the statistical robustness of the data and accounts for both inter-experimental and intra-experimental variations. Data analysis, including ANOVA, standard curve plotting, and the correct interpretation and comparison of different detection methods, was performed using GraphPad Prism (version 7). A p < 0.05 was considered statistically significant (*p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001).
Validation of protocol
This protocol has been used and validated in the following research article:
Cao et al. [1]. A DNAzyme-enhanced nonlinear hybridization chain reaction for sensitive detection of microRNA. Journal of Biological Chemistry (Figure 4, panel A/B; Figure 5, panel C/D).
In Table 6, the detection performance of this platform is compared with other miRNA detection methods.
Table 6. Comparison of the proposed method with other microRNA detection methods
Methods Target miRNA Limit of detection Linear detection range Characteristic Ref.
qPCR miRNAs 25 copies 102–106 copies Sensitive, specific, bulky equipment and trained personnel requirement [20]
Northern blot miR-16
miR-18a
miR-30a
25 amol n/a Multiplexing, time consuming, and semiquantitative [21]
Microarray miR let 7a 0.080 pM 0.2 pM–1.50 nM Low specificity, high-throughput, selective, and sensitive [22]
RCA miR let 7a 10 fM 25 fM–2.5 pM Robustness, specificity, high sensitivity, and contains enzymes [23]
SDA miRNAs 16 zM 1.0 pM–0.1 zM Wide linear range, sensitive, and contains enzymes [24]
Catalytic hairpin assembly miR-21 1 pM 1 pM–100 nM Enzyme-free, stable, low cost, and sensitive [25]
LAMP miR let 7a 1.0 aM 1.0 aM–1.0 pM Specific, sensitive, fast, and contains enzymes [26]
bHCR-DNAzyme miR-21 0.004 nM 0.01 nM–0.2 nM Fast, convenient, low cost, specific, and enzyme-free This work
Note: LAMP, loop-mediated isothermal amplification; qPCR, quantitative polymerase chain reaction; RCA, rolling circle amplification; SDA, strand-displacement amplification.
General notes and troubleshooting
General notes
When measuring the fluorescence values of the reaction system using a fluorescence spectrophotometer, ensure that the parameter settings remain consistent each time to avoid changes.
Handle systems containing the H2 probe with care to avoid light exposure. The H2 probe is labeled with a fluorescent group, so preventing it from absorbing specific wavelengths of light that could cause excitation is crucial for maintaining the stability of experimental results.
Perform hairpin formation for probes H1/H2/H3/H4 before use but be cautious not to prepare excessive amounts at once. Repeated freeze-thaw cycles can lead to degradation of the hairpin structures, potentially compromising the stability of experimental results.
Troubleshooting
Problem 1: High background signal in images when validating bHCR-DNAzyme using non-denaturing polyacrylamide gel electrophoresis (PAGE).
Possible cause: Poor quality of the electrophoresis buffer or insufficient washing time after staining.
Solution: Replace with fresh electrophoresis buffer and increase the number of washes with buffer after staining.
Problem 2: The measured fluorescence intensity value exceeds the maximum limit of the y-axis display.
Possible cause: The concentration of hairpin in the reaction system is too high, and the measured fluorescence intensity value exceeds the maximum value of the visible fluorescence intensity under the current parameter.
Solution: Measure after diluting the reaction system or change measurement parameters, such as reducing the voltage and reducing the slit width.
Acknowledgments
This work was supported by the National Natural Science Foundation of China [C0602-32270609]; the Hunan Provincial Natural Science Foundation of China [2021JJ30916]; and the Hunan Provincial Natural Science Foundation for Distinguished Young Scholars [2022JJ10091]. This protocol was described and validated in the following previous work: Cao et al. [1], J Biol Chem. 2023 Jun;299(6):104751.
Competing interests
The authors declare no competing interests.
Ethical considerations
Tumor tissue and PT tissue obtained from patients with hepatocellular carcinoma from the Third Xiangya Hospital in Central South University, Approval No: 2023009; all experiments were conducted following the Declaration of Helsinki and good clinical practice guidelines.
References
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Article Information
Publication history
Received: Aug 27, 2024
Accepted: Oct 10, 2024
Available online: Nov 4, 2024
Published: Dec 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Total Internal Reflection Fluorescence (TIRF) Single-Molecule Assay to Analyze the Motility of Kinesin
TK Tomoki Kita
SN Shinsuke Niwa
Published: Vol 14, Iss 24, Dec 20, 2024
DOI: 10.21769/BioProtoc.5135 Views: 463
Reviewed by: Neha NandwaniDjamel Eddine ChafaiXi Feng
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Original Research Article:
The authors used this protocol in eLIFE Jan 2024
Abstract
The motile parameters of kinesin superfamily proteins are fundamental to intracellular transport. Single-molecule motility assays using total internal reflection fluorescence (TIRF) microscopy are a gold standard technique for measuring the motile parameters of kinesin motors. With this technique, one can evaluate the velocity, run length, and binding frequency of kinesins on microtubules by directly observing their motility. This protocol provides a comprehensive procedure for single molecule assays of kinesins, including the preparation of labeled microtubules, the measurement of kinesin motility via TIRF microscopy, and the quantification of kinesin motor parameters.
Key features
• Analysis of the motility of kinesin superfamily proteins using TIRF microscopy.
• In vitro reconstitution using purified microtubules and motors.
• Direct measurement of motile parameters of kinesins.
Keywords: Kinesin Microtubules Single-molecule assay KIF1A UNC-104
Background
Intracellular transport is essential for many cellular processes. Kinesin is an ATP-dependent molecular motor that moves along microtubules and facilitates anterograde transport [1]. Organisms have many kinesins, and each kinesin has motile properties specific to its function [2]. Fluorescence microscopy allows us to observe kinesins or their cargo labeled with fluorescent proteins. However, background fluorescence from outside the focal plane can hinder the observation of single fluorophores and the examination of kinesin motility at a single-molecule resolution. In TIRF microscopy, the laser reflects between the coverslip and solution to form an evanescent field that excites the fluorescent protein on the motor [3]. This excitation is restricted to a region typically less than 100 nm in thickness, allowing us to track kinesins with single fluorophores [3].
Using TIRF microscopy, single-molecule parameters of kinesins, such as velocity and run length, have been well studied [4,5]. Disease-associated motile defects of kinesins can be directly measured by this method [6–9]. Additionally, the assay can analyze the extent of kinesin activation [10,11]. Most kinesins are inactivated by autoinhibition to avoid binding to microtubules without cargo [12]. Upon release from autoinhibition, the motor’s landing rate on microtubules increases dramatically, enabling us to observe their activated movements along microtubules [10,11].
In this protocol, we present three methods for single-molecule assays. The first involves modifying microtubules to stabilize them on glass surfaces using biotin-streptavidin interactions and visualize them with fluorescence. The second focuses on observing the motility of individual kinesin molecules, and the third covers their subsequent analysis including velocity, run length, and landing rate. Our comprehensive protocol is well-suited for researchers entering the field of kinesin molecular motors.
Materials and reagents
Biological materials
Porcine tubulin; tubulin was purified from porcine brain, as described [13].
Kinesin motor of interest; in this protocol, KIF1A(1-393)LZ::mScarletI, UNC-104(1-653)::sfGFP and UNC-104(1-653)(E412K)::sfGFP are used as examples for TIRF assays. Protein was produced as previously described using LOBSTR-BL21(DE3)-RIL E. coli (Kerafast, catalog number: EC1002) in the case of KIF1A(1-393)LZ::mScarlet [8] and Sf9 insect cells in the case of UNC-104(1-653)::sfGFP and UNC-104(1-653)(E412K)::sfGFP [4].
Reagents
Piperazine-1,4-bis(2-ethanesulfonic acid) (PIPES) (FUJIFILM Wako Pure Chemical Corporation, catalog number: 345-02225)
Magnesium chloride (MgCl2) (Sigma-Aldrich, catalog number: M2393)
Ethylene glycol tetraacetic acid (EGTA) (Nacarai Tesque, catalog number: 15214-92)
Potassium hydroxide (KOH) (Nacarai Tesque, catalog number: 28616-45)
2-[4-(2-Hydroxyethyl)-1-piperazinyl]ethanesulfonic acid (HEPES) (Nacarai Tesque, catalog number: 17514-15)
Sodium hydroxide (NaOH) (FUJIFILM Wako Pure Chemical Corporation, catalog number: 191-18875)
Glycerol (FUJIFILM Wako Pure Chemical Corporation, catalog number: 072-00621)
L-Glutamic acid potassium salt (K-Glutamate) (Sigma-Aldrich, catalog number: G1501)
Paclitaxel (Taxol) (Sigma-Aldrich, catalog number: T7402)
Dimethyl sulfoxide (DMSO) (Nacarai Tesque, catalog number: 09659-14)
AFDye 647 NHS (Vector Laboratories, catalog number: FP-1121)
Biotin-PEG2-NHS (Tokyo Chemical Industry Co., Ltd., catalog number: B6097)
Guanosine 5’-triphosphate (GTP) (Nacarai Tesque, catalog number: 17450-61)
PLL-PEG biotin [SuSoS AG, product name: PLL(20)-g[3.5]- PEG(2)/PEG(3.4)- biotin(50%)]
Streptavidin (FUJIFILM Wako Pure Chemical Corporation, catalog number: 194-17863)
Potassium acetate (KCH3COO) (FUJIFILM Wako Pure Chemical Corporation, catalog number: 160-03175)
Pluronic F-127 (Sigma-Aldrich, catalog number: P2443)
Magnesium Acetate [Mg(CH3COO)2] (FUJIFILM Wako Pure Chemical Corporation, catalog number: 135-10011)
κ-Casein (Sigma-Aldrich, catalog number: C0406)
Bovine serum albumin (BSA) (Nacarai Tesque, catalog number: 01863-06)
Biotin-BSA (Thermo Fisher Scientific, catalog number: 29130)
Adenosine 5’-triphosphate (ATP) (Nacarai Tesque, catalog number: 01072-82)
Protocatechuic acid (PCA) (Nacarai Tesque, catalog number: 08521-24)
Protocatechuate-3,4-dioxygenase (PCD) (TOYOBO, catalog number: PCO-302)
Trolox (FUJIFILM Wako Pure Chemical Corporation, catalog number: 202-17891)
Methanol (FUJIFILM Wako Pure Chemical Corporation, catalog number: 134-01833)
Ethanol (FUJIFILM Wako Pure Chemical Corporation, catalog number: 057-00541)
Hydrogen chloride (HCl) (FUJIFILM Wako Pure Chemical Corporation, catalog number: 087-01076)
Sucrose (Nacarai Tesque, catalog number: 30403-55)
Vaseline (Nacarai Tesque, catalog number: 36202-05)
Lanolin (FUJIFILM Wako Pure Chemical Corporation, catalog number: 128-00115)
Paraffin (Nacarai Tesque, catalog number: 26023-65)
Solutions
BRB80 (see Recipes)
High-pH cushion (see Recipes)
Labeling buffer (see Recipes)
Quench buffer (see Recipes)
Low-pH cushion (see Recipes)
1 mM Taxol (see Recipes)
Sucrose cushion (see Recipes)
Assay buffer (see Recipes)
100 mM Trolox (see Recipes)
VALAP (see Recipes)
Recipes
BRB80
80 mM PIPES (adjust the pH to 6.8 with KOH)
1 mM MgCl2
1 mM EGTA
High-pH cushion
0.1 M HEPES (adjust the pH to 8.6 with NaOH)
1 mM MgCl2
1 mM EGTA
60% glycerol
Labeling buffer
0.1 M HEPES (adjust the pH to 8.6 with NaOH)
1 mM MgCl2
1 mM EGTA
40% glycerol
Quench buffer
2× BRB80
100 mM K-Glutamate (adjust the pH to 7.0 with KOH)
40% glycerol
Low-pH cushion
1× BRB80
60% glycerol
1 mM Taxol
Dissolve Taxol in DMSO.
Sucrose cushion
Dissolve 3 g of sucrose in BRB80 with 20 μM Taxol and make up the volume to 10 mL. Rotate the solution at 35 °C overnight.
Assay buffer
180 μL of 0.5 M HEPES (adjust the pH to 7.4 with KOH)
200 μL of 50% glycerol
50 μL of 1 M KCH3COO
50 μL of 10% Pluronic F-127
20 μL of 100 mM Mg(CH3COO)2
20 μL of 50 mM EGTA
20 µL of 10 mg/mL κ-Casein
20 μL of 50 mg/mL BSA
20 μL of 5 mg/mL biotin-BSA
1 μL of 1 mM Taxol
100 mM Trolox
Mix 430 μL of methanol and 345 μL of 1 M NaOH. Dissolve 100 mg of Trolox in the solution and make up the volume to 4 mL with ddH2O. Rotate the solution at 25 °C overnight. Pass the solution through a 0.22 μm filter.
VALAP
50 g of Vaseline
50 g of lanolin
50 g of paraffin
Laboratory supplies
Pipette tips
2–20 μL [Thermo Scientific (QSP), catalog number: TLR102RL-Q]
10–100 μL [Thermo Scientific (QSP), catalog number: 110RL-NEW]
100–1,000 μL [Thermo Scientific (QSP), catalog number: T112XLRL-Q]
1.5 mL microtubes (WATSON, catalog number: 131-415C)
Thick wall polycarbonate tubes
0.2 mL (Beckman Coulter, catalog number: 343775)
1 mL (Beckman Coulter, catalog number: 343778)
3.5 mL (Beckman Coulter, catalog number: 349622)
0.22 μm filter (MERCK, catalog number: UFC30GV00)
Coverslips (THORLABS, catalog number: CG15CH)
Microscope slides (MATSUNAMI, catalog number: S1225)
Double-sided tape (NICHIBAN, catalog number: NW-25)
Equipment
Micropipette
0.5–10 μL (Eppendorf, catalog number: 4924000029)
2–20 μL (Eppendorf, catalog number: 4924000037)
20–200 μL (Eppendorf, catalog number: 4924000061)
100–1,000 μL (Eppendorf, catalog number: 4924000088)
Ultracentrifuge (Beckman Coulter, model: Optima TLX Ultracentrifuge)
Centrifuge rotor (Beckman Coulter, model: TLA-100, TLA120.2, TLA100.3)
Water bath (TAITEC, model: CTU-mini, catalog number: 0063288-000)
Water bath with sonicator (AS ONE, model: ASU-M)
TIRF setup
Fluorescence microscope (Nikon Instruments, model: ECLIPSE Ti2-E)
Objective lens (Nikon Instruments, model: CFI Apochromat TIRF 100XC Oil)
Glass heater (TOKAI HIT, model: TPiD-SQH26-LH)
Camera (Oxford Instruments, model: Andor iXion life 897)
Laser (Nikon Instruments, model: Ti2-LAPP illumination system)
System software (Nikon Instruments, model: NIS-Elements AR software version 5.2)
Software and datasets
ImageJ Fiji [14]
Procedure
Labeling tubulin with biotin or fluorescent dye
Prepare biotin-labeled tubulin and fluorescently labeled tubulin to stabilize microtubules on a streptavidin-coated glass surface and visualize them [15].
Make 7 mL of 3 mg/mL tubulin solution by diluting the 7–10 mg/mL purified tubulin solution [13] with BRB80 on ice.
Add 35 μL of 1 M MgCl2, 100 μL of 100 mM GTP, and 3.5 mL of glycerol to the solution and gently mix. Incubate the solution in a water bath at 37 °C for 60 min.
Layer 1.75 mL of the solution on 1.4 mL of high-pH cushion in six 3.5 mL thick-wall polycarbonate tubes. Centrifuge at 80,000× g for 35 min at 35 °C in the TLA100.3 rotor. High-pH cushion retains non-polymerized tubulin in the supernatant and raises the buffer pH sufficiently for the labeling reaction.
Aspirate the supernatant slowly and wash the interface of the cushion with 1 mL of 37 °C warm labeling buffer twice. Remove the cushion and thoroughly resuspend the pellet in the first tube with 500 μL of 37 °C warm labeling buffer, then sequentially transfer and resuspend in the next tubes up to the sixth tube to combine all pellets into 500 μL of solution.
Add 1 mg of AFDye 647 NHS or Biotin-PEG2-NHS dissolved in DMSO to the solution and vortex it quickly. Incubate the solution in a water bath at 37 °C for 10 min and lightly vortex every 2 min. Add 530 μL of warm (37 °C) quench buffer to the solution and lightly vortex to stop the labeling reaction. Incubate the solution in a water bath at 37 °C for 5 min.
Layer 1 mL of the solution on 1.5 mL of low-pH cushion in a 3.5 mL thick-wall polycarbonate tube. Centrifuge at 80,000× g for 20 min at 35 °C in the TLA100.3 rotor. Low-pH cushion retains the free dye in the supernatant.
Aspirate the supernatant slowly and wash the interface of the cushion with warm (37 °C) BRB80 twice. Remove the cushion and thoroughly resuspend the pellet with 500 μL of cold (4 °C) BRB80 on ice.
Transfer the solution into a 1 mL thick-wall polycarbonate tube and centrifuge at 80,000× g for 10 min at 4 °C in the TLA120.2 rotor. In this procedure, non-depolymerized tubulin resulting from the labeling process is precipitated and removed.
Take out the supernatant and add 130 μL of 5× BRB80, 2 μL of 1 M MgCl2, and 5 μL of 100 mM GTP on ice.
Add 330 μL of glycerol to the solution and gently mix. Incubate the solution in a water bath at 37 °C for 30 min.
Layer 1 mL of the solution on 1.5 mL of low-pH cushion in a 3.5 mL thick-wall polycarbonate tube. Centrifuge at 80,000× g for 35 min at 35 °C in the TLA100.3 rotor. Low-pH cushion retains the non-polymerized tubulin in the supernatant.
Aspirate the supernatant slowly and wash the interface of the cushion with warm (37 °C) BRB80 twice. Remove the cushion and wash the interface of the pellet with warm (37 °C) BRB80 twice. Resuspend the pellet with 200 μL of cold (4 °C) BRB80 and incubate on ice for 20 min.
Centrifuge at 80,000× g for 10 min at 4 °C in the TLA100 rotor. Finally, non-depolymerized tubulin is precipitated and removed.
Measure the concentration of labeled tubulins using SDS-PAGE. As a control, use unlabeled tubulins with a known concentration.
Aliquot the supernatant in 3 μL portions, freeze in liquid nitrogen, and store at -80 °C.
Microtubule polymerization
To make 25 μL of elongation mix:
20 μL of unlabeled tubulin (3 mg/mL)
2 μL of biotin-labeled tubulin (3 mg/mL)
2 μL of fluorescently labeled tubulin (3 mg/mL)
1 μL of 100 mM GTP
The concentrations of biotin-labeled and fluorescently labeled tubulin should be approximately 1/10 to 1/40 of the unlabeled tubulin. In practice, their concentrations should be minimized as much as possible, as excessive labeling can hinder motor movement. It is sufficient for the overall microtubule fluorescence on the glass to be just barely detectable under the microscope.
Incubate the mix in a water bath at 37 °C for more than 20 min. Microtubules that are approximately 10–20 μm in length are polymerized after 20 min of incubation. Incubating for around 45 min results in microtubules approximately 50 μm long. However, microtubules longer than 50 μm are generally unnecessary, as they often extend beyond the field of view in TIRF microscopy.
Add 25 μL of 40 μM Taxol to the mix and incubate at 37 °C for more than 15 min.
Layer the mix on 150 μL of sucrose cushion in a 0.2 mL thick-wall polycarbonate tube. Centrifuge at 50,000× g for 10 min at 35 °C in the TLA-100 rotor. Sucrose cushion retains the non-polymerized tubulin in the supernatant.
Aspirate the supernatant slowly and wash the interface of the cushion with warm (37 °C) BRB80 twice. Remove the cushion and thoroughly resuspend the pellet with 50 μL of BRB80 with 20 μM Taxol.
Store the microtubules at room temperature and protect from light.
TIRF single-molecule assay
Wash glass chamber.
Acid wash the coverslips with HCl for 24 h and rinse with ddH2O.
Fill the container with ddH2O and sonicate in a water bath at 42 kHz three times for 30 min each.
Fill the container with 50%, 70%, and 95% ethanol sequentially, and sonicate in a water bath at 42 kHz for 30 min for each concentration.
Fill the container with 95% ethanol and store at room temperature.
Make a glass chamber.
Burn both sides of the coverslip with a gas burner to volatilize the ethanol (Figure 1A).
Attach the coverslip to a glass slide using two pieces of double-sided tape at the top and bottom, creating open slits on both sides (Figure 1B). The channel width is approximately 3 mm. If the observation time in one channel is short enough to prevent evaporation in the second channel (approximately 1–3 min), two channels can be prepared on a single coverslip to quickly examine two conditions (Figure 1C).
Figure 1. Illustration of glass slide chamber. (A) Burning both sides of a coverslip with a gas burner. (B) Making glass slide chamber using double-sided tape. (C) Preparing two channels for examining and comparing two conditions.
Flow in 10 μL of 0.5 mg/mL PLL-PEG-biotin into the chamber and let it sit for 5 min. PLL-PEG coats the glass surface and prevents the non-specific binding of proteins.
Flow in 10 μL of 0.5 mg/mL streptavidin into the chamber and let it sit for 2 min.
Wash out the unbound streptavidin with 2 × 20 μL of BRB80.
Flow in 10 μL of labeled microtubules at a 1/50 dilution (in BRB80 with 20 μM Taxol) into the chamber and let it sit for more than 5 min.
Wash out the unbound microtubules with 2 × 20 μL of assay buffer.
To make 50 μL of observation mix:
29 μL of assay buffer
16 μL of ddH2O
1 μL of 10 pM to 10 nM purified kinesin (depending on the type of kinesin)
1 μL of 100 mM ATP
1 μL of 100 mM PCA
1 μL of 2.5 mM PCD
1 μL of 100 mM Trolox
PCA, PCD, and Trolox serve as an oxygen scavenging system.
Flow in 10 μL of the observation mix into the chamber.
Observe the single-molecule motility of motor proteins under the TIRF microscope at room temperature (24 ± 1 °C). In this protocol, 15 mW 488 nm, 561 nm, and 647 nm excitation lasers are used. Videos 1–3 are examples of TIRF single-molecule assays of KIF1A/UNC-104 (kinesin-3). Video 1 shows the motility of artificially dimerized truncated KIF1A using the leucine zipper domain [KIF1A(1-393)LZ]. The red fluorescent protein mScarlet I was attached to its C-terminal. In the first frame, an image of microtubules was taken using 640 nm excitation (laser strength: 9%). In the remaining frames, images of KIF1A(1-393)LZ were taken using 561 nm excitation (laser strength: 5%) at 10 frames per second (fps) and saved in the nd2 file format. In Video 1, images from two color channels were merged. Videos 2 and 3 show the motility of truncated UNC-104 [UNC-104(1-653)] and its mutant [UNC-104(1-653)(E412K)], respectively. The green fluorescent protein sfGFP was attached to their C-terminals. In the first frame, an image of microtubules was taken using 640 nm excitation (laser strength: 9%). In the remaining frames, images of UNC-104 were taken using 488 nm excitation (laser strength: 5%) at 10 fps and saved in the nd2 file format. In Videos 2 and 3, images from two color channels were merged.
Video 1. Example of total internal reflection fluorescence (TIRF) single-molecule assay of KIF1A(1-393)LZ labeled by mScarlet I. Video is played at 5× speed. Scale bar shows 10 μm.
Video 2. Example of total internal reflection fluorescence (TIRF) single-molecule assay of UNC-104(1-653) labeled by sfGFP. Video is played at 5× speed. Scale bar shows 10 μm.
Video 3. Example of total internal reflection fluorescence (TIRF) single-molecule assay of UNC-104(1-653)(E412K) labeled by sfGFP. Video is played at 5× speed. Scale bar shows 10 μm.
Data analysis
Figure 2 illustrates the principle of a TIRF single-molecule assay. Fluorescent microtubules are fixed onto a PLL-PEG biotin and streptavidin-coated coverslip. An aliquot of kinesins labeled with fluorescent proteins is introduced into the chamber. The movements of the fluorescently labeled kinesins are observed as they move continuously in a unidirectional manner.
Figure 2. Illustration of a single-molecule assay. The coverslip is coated with PLL-PEG-biotin followed by streptavidin. Microtubules are then flowed into the chamber and fixed onto the coverslip. Fluorescently labeled kinesins are introduced into the chamber. Kinesin motility is observed under a total internal reflection fluorescence (TIRF) microscope.
ImageJ Fiji was used to create kymographs for the analysis of kinesin. Import the nd2 file into ImageJ Fiji. Use the Segmented Line tool to draw a line along the microtubule in the 640 nm channel. Create a kymograph using the KymographBuilder plugin. Figure 3 illustrates the principle of kymograph analysis. Figure 3A shows a series of images taken at different time points in Video 1, with kinesins moving from left to right as time progresses. Similarly, the kymograph is a two-dimensional representation, where the x-axis represents time, and the y-axis represents the spatial position along the selected microtubule (Figure 3B). In the kymograph, moving objects appear as diagonal lines (Figure 3B).
Figure 3. Unidirectional movements of kinesins and its kymograph. (A) Sequential frames showing representative single-kinesin movements from Video 1. A scale bar shows 5 μm. (B) The kymograph of (A). Horizontal and vertical bars show 5 μm and 5 s, respectively.
We demonstrate the analysis of the TIRF single-molecule assay of UNC-104. UNC-104 is a kinesin-3 family protein that transports synaptic vesicle precursors in C. elegans axons. UNC-104(1-653), a cargo-binding domain deletion mutant, is strongly autoinhibited and rarely bound to microtubules (Video 2). In contrast, UNC-104(1-653)(E412K), an autoinhibition-disrupted mutant, is frequently bound to microtubules (Video 3). To quantify its extent of activation, we analyzed velocity, run length, and landing rate from their kymographs (Figure 4A). The slope of the diagonal lines indicates the velocity of the movement: steeper slopes represent faster movement, while shallower slopes indicate slower movement. The distance along the x-axis between the attachment and detachment points of motors on the microtubule represents the run length. The landing rate is calculated by dividing the number of motor attachments on the microtubule by the observation time and length of the microtubule. The run length and landing rate of UNC-104(1-653)(E412K) significantly increased compared to UNC-104(1-653), but their velocities remained the same [4] (Figure 4B–D).
Figure 4. Representative results. (A) Representative kymographs of UNC-104(1-653) and UNC-104(1-653)(E412K) from Video 2 and Video 3, respectively. Horizontal and vertical bars show 10 μm and 10 s, respectively. (B) The velocities of UNC-104(1-653) and UNC-104(1-653)(E412K) are plotted as violin graphs. Green bars represent mean ± SD. Student’s t-test, ns, p = 0.7 and statistically not significant. (C) The run lengths of UNC-104(1-653) and UNC-104(1-653)(E412K) are plotted as violin graphs. Green bars represent the median value and interquartile range. Mann-Whitney U test. ****, p < 0.0001. (D) The landing rates of UNC-104(1-653) and UNC-104(1-653)(E412K) are plotted as violin graphs. Green bars represent the median value. Mann-Whitney U test. ****, p < 0.0001. The values in (B)–(D) are identical to those reported in Kita et al. [4]; see the reference for further details.
Notes:
Polymerized microtubules can be stored at room temperature for 2–3 weeks. Microtubules are depolymerized at low temperatures and should not be refrigerated.
Kinesin proteins must be stored at -80 °C.
Single-molecule assays for kinesins are typically performed at room temperature (24 ± 1 °C). The chamber temperature can be regulated using a glass heater. The temperature dependency varies depending on the type of motor [16].
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Kita et al. [4]. Comparative analysis of two Caenorhabditis elegans kinesins KLP-6 and UNC-104 reveals a common and distinct activation mechanism in kinesin-3 eLife (Figure 1D, Figure 2, Figure 3C–E, Figure 4D–F, Figure 5D–G, and Figure 7D–G).]
General notes and troubleshooting
General notes
This protocol can be applied to analyze other kinesin superfamily proteins and dynein motor proteins from different species. Some motor proteins may be very slow. In that case, one needs to adjust the frame rate. 100 ms/frame for 1 min is the starting point. We sometimes observe very slow motors at 2 s/frame for 10 min. To prevent evaporation in the channels, it is recommended to seal the channels with VALAP during observation.
Troubleshooting
Problem 1: Microtubules are short.
Possible causes: Excessive pipetting and/or insufficient incubation time.
Solutions: Reduce pipetting or use gentle tapping instead. Extend the incubation time at 37 °C to 45 min.
Problem 2: Microtubules are not stabilized on the glass and/or are not visualized with fluorescence.
Possible causes: Insufficient biotin-labeled tubulin and/or fluorescently labeled tubulin.
Solutions: Increase the concentration of biotin-labeled and/or fluorescently labeled tubulin during polymerization. However, note that excessive labeling can impede kinesin movement. It is sufficient for the overall microtubule fluorescence on the glass to be just barely detectable under the microscope.
Problem 3: Kinesins move very slowly or do not move.
Possible causes: Excessive labeling of microtubules.
Solutions: Reduce the concentration of biotin-labeled and/or fluorescently labeled tubulin during polymerization. KIF1A(1-393)LZ can serve as a reliable positive control to verify the conditions. In our setup, it moves at approximately 1.5 μm/s along microtubules.
Acknowledgments
We would like to thank the members of the Niwa lab (Tohoku University). T.K. was supported by JSPS KAKENHI (23KJ0168). S.N. was supported by JSPS KAKENHI (23H02472) and the Naito foundation. This method was originally described and validated in our paper [4].
Competing interests
The authors have no competing interests.
References
Vale, R. D., Reese, T. S. and Sheetz, M. P. (1985). Identification of a novel force-generating protein, kinesin, involved in microtubule-based motility. Cell. 42(1): 39–50.
Hirokawa, N., Noda, Y., Tanaka, Y. and Niwa, S. (2009). Kinesin superfamily motor proteins and intracellular transport. Nat Rev Mol Cell Biol. 10(10): 682–696.
Vale, R. D., Funatsu, T., Pierce, D. W., Romberg, L., Harada, Y. and Yanagida, T. (1996). Direct observation of single kinesin molecules moving along microtubules. Nature. 380(6573): 451–453.
Kita, T., Chiba, K., Wang, J., Nakagawa, A. and Niwa, S. (2024). Comparative analysis of two Caenorhabditis elegans kinesins KLP-6 and UNC-104 reveals a common and distinct activation mechanism in kinesin-3. eLife. 12: e89040.
Fan, X. and McKenney, R. J. (2023). Control of motor landing and processivity by the CAP-Gly domain in the KIF13B tail. Nat Commun. 14(1): 4715.
Lam, A. J., Rao, L., Anazawa, Y., Okada, K., Chiba, K., Dacy, M., Niwa, S., Gennerich, A., Nowakowski, D. W., McKenney, R. J., et al. (2021). A highly conserved 310 helix within the kinesin motor domain is critical for kinesin function and human health. Sci Adv. 7(18): eabf1002.
Boyle, L., Rao, L., Kaur, S., Fan, X., Mebane, C., Hamm, L., Thornton, A., Ahrendsen, J. T., Anderson, M. P., Christodoulou, J., et al. (2021). Genotype and defects in microtubule-based motility correlate with clinical severity in KIF1A-associated neurological disorder. Hum Genet Genomics Adv. 2(2): 100026.
Anazawa, Y., Kita, T., Iguchi, R., Hayashi, K. and Niwa, S. (2022). De novo mutations in KIF1A-associated neuronal disorder (KAND) dominant-negatively inhibit motor activity and axonal transport of synaptic vesicle precursors. Proc Natl Acad Sci USA. 119(32): e2113795119.
Chiba, K., Takahashi, H., Chen, M., Obinata, H., Arai, S., Hashimoto, K., Oda, T., McKenney, R. J. and Niwa, S. (2019). Disease-associated mutations hyperactivate KIF1A motility and anterograde axonal transport of synaptic vesicle precursors. Proc Natl Acad Sci USA. 116(37): 18429–18434.
Monroy, B. Y., Tan, T. C., Oclaman, J. M., Han, J. S., Simó, S., Niwa, S., Nowakowski, D. W., McKenney, R. J. and Ori-McKenney, K. M. (2020). A Combinatorial MAP Code Dictates Polarized Microtubule Transport. Dev Cell. 53(1): 60–72.e4.
Chiba, K., Ori-McKenney, K. M., Niwa, S. and McKenney, R. J. (2022). Synergistic autoinhibition and activation mechanisms control kinesin-1 motor activity. Cell Rep. 39(9): 110900.
Chiba, K. and Niwa, S. (2024). Autoinhibition and activation of kinesin-1 and their involvement in amyotrophic lateral sclerosis. Curr Opin Cell Biol. 86: 102301.
Castoldi, M. and Popov, A. V. (2003). Purification of brain tubulin through two cycles of polymerization–depolymerization in a high-molarity buffer. Protein Expression Purif. 32(1): 83–88.
Schindelin, J., Arganda-Carreras, I., Frise, E., Kaynig, V., Longair, M., Pietzsch, T., Preibisch, S., Rueden, C., Saalfeld, S., Schmid, B., et al. (2012). Fiji: an open-source platform for biological-image analysis. Nat Methods. 9(7): 676–682.
Al-Bassam, J. (2014). Chapter Eight - Reconstituting Dynamic Microtubule Polymerization Regulation by TOG Domain Proteins. In: Vale, R. D. (Ed.). Methods in Enzymology, Academic Press, volume 540 of Reconstituting the Cytoskeleton, 131–148.
Kushwaha, V. S. and Peterman, E. J. (2020). The temperature dependence of kinesin motor-protein mechanochemistry. Biochem Biophys Res Commun. 529(3): 812–818.
Article Information
Publication history
Received: Aug 6, 2024
Accepted: Oct 5, 2024
Available online: Oct 29, 2024
Published: Dec 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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5,136 | https://bio-protocol.org/en/bpdetail?id=5136&type=0 | # Bio-Protocol Content
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Egg Microinjection for the Ladybird Beetle Harmonia axyridis
TN Taro Nakamura
YM Yuji Matsuoka
TN Teruyuki Niimi
Published: Vol 14, Iss 24, Dec 20, 2024
DOI: 10.21769/BioProtoc.5136 Views: 306
Reviewed by: Alberto RissoneKai Yuan Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in PLOS ONE Jun 2014
Abstract
In this paper, we present a detailed protocol for microinjecting DNA, RNA, or protein solutions into fertilized eggs of the multicolored Asian ladybird beetle, Harmonia axyridis, under a stereomicroscope equipped with an injection apparatus. H. axyridis is an emerging model organism for studying various biological fields, showing intraspecific polymorphisms exhibiting highly diverse color patterns on the elytra. Here, we describe how to rear ladybird beetles in a laboratory and obtain fertilized eggs for microinjection experiments. We also provide a constant fluid flow injection method, which enhances the efficiency of microinjection and improves throughput. Our step-by-step protocol is applicable to generating transgenic or genome-edited ladybird beetles, facilitating functional genetics in H. axyridis; the microinjection method should be applicable to other insect eggs.
Key features
• Egg microinjection for the multicolored Asian ladybird beetle Harmonia axyridis under a stereomicroscope with injection equipment.
• Detailed procedures for post-injection care, including rearing Harmonia axyridis.
• Step-by-step guide for the efficient collection of Harmonia axyridis eggs.
Keywords: Egg microinjection Ladybird beetles Harmonia axyridis Gene modification
Graphical overview
Background
Ladybird beetles belong to the order Coleoptera and encompass approximately 6,000 species worldwide, with around 190 species being described in Japan [1,2]. These beetles display diverse color patterns on their elytra, primarily black and red, which are not only species-specific but also display a high degree of diversity within species (intraspecific polymorphism) (reviewed in Ando and Niimi [3]). Of ladybird beetles, Harmonia axyridis has become an emerging model organism in diverse biological fields, such as color patterning [4,5], population genetics of intraspecific polymorphism [6,7], biological insecticides [8], and phenotypic plasticity [9]. H. axyridis possesses desirable characteristics as an experimental animal, such as the availability of an artificial diet, long-term storage of adults at low temperatures (up to 2 years and 8 months; personal observation), commercially available adults, high fecundity, easy access to fertilized eggs, and ease of sexing in adults based on morphology and at any stage by PCR-based techniques [10]. The ladybird beetle's lifecycle is relatively short—approximately 3 days for embryogenesis, around 11 days for larval development through three molts, 4.5 days for pupal development, and a lifespan of 2–3 months for adults (Figure 1). Furthermore, modern experimental techniques, including high-quality genome data [11,12], cryopreservation of the gonad [13], and numerous useful transgenic lines and mutant strains [14–16] including flightless mutants [17], and modern functional genomics, including RNAi, transgenics, and genome editing, have been developed for this species [4,14,15,18–20].
Figure 1. Life stages of Harmonia axyridis. The developmental period for each stage under laboratory conditions at 25 °C is indicated at the bottom. The period shown for adults represents the time during which they lay eggs after mating. It takes approximately one week after eclosion for adults to reach sexual maturity. Scale bar: 1 mm.
Fertilized egg microinjection is an essential approach for functional genomics including transgenesis and genome editing, being the most commonly used method in insects. While there are several methods for delivering reagents into eggs—including electroporation [21–23] and particle guns [24–26], which enable simultaneous treatment of multiple eggs—microinjection is the most widely used method for introducing nucleic acids, proteins, and chemicals into insect eggs. This method, however, has problems such as frequent clogging due to the backflow of egg contents, which prevents efficient injection. To overcome this problem, we developed an injection method using constant fluid flow (i.e., constant pressure during injection). This method not only prevents needle clogging but also contributes to increased injection throughput because it allows the delivery of the injection solution without egg-by-egg manual injection. However, this microinjection system is not able to control a precise injection volume, but the injection volume can be controlled by adjusting the time the needle is kept inside the egg. Our injection system is performed under a stereomicroscope with a micromanipulator and an injection needle holder, making the construction of an injection system more affordable than systems with compound microscopes. This paper describes the methodology of microinjection into the multicolored Asian ladybird beetle embryos using constant fluid flow injection with a stereomicroscope.
Materials and reagents
Biological materials
Harmonia axyridis (young-adult female and male beetles). Ladybird beetles are commercially available as biopesticides in Japan (Tentop, Agrisect, Japan) and possibly other countries. We used established lab culture lines once they had been collected in Aichi, Japan from the outside environment in the spring.
Pea aphid, Acyrthosiphon pisum (feeding source, important for high egg productivity)
Note: Live pea aphids are not commercially available. Pea aphids are found all over the world and can be self-fertilized. Aphids can be maintained on broad bean plants and are easily reared as they are asexual to ensure a steady food supply for the beetles.
Laboratory supplies
Micro-loading tip (Eppendorf, catalog number: 5242956003)
Paper towel (laid on the bottom of the rearing case)
Double-sided tape (Nitoms, catalog number: J0820 or equivalent)
Slide glass (Matsunami, catalog number: S1111)
Artist’s brush (00 number painting brush)
Glass capillary (NARISHIGE, catalog number: GD-1)
Equipment
FemtoJet 4i (Eppendorf, catalog number: 5252000021)
Dissection/stereomicroscope (Zeiss or equivalent)
Micromanipulator (Narishige, catalog number: MMO-202ND)
Micromanipulator stand (Narishige, catalog number: GJ-1)
Micromanipulator stand base (Narishige, catalog number: IP)
Needle holder (Narishige, catalog number: HI-7)
Micropipette puller (Sutter Instruments, catalog number: P-1000)
Box filament (FB245B, 2.5 mm × 4.5 mm)
Microcentrifuge (TOMY, model: MX-307)
Pipetman [GILSON, catalog numbers: F123602 (P1000), F123601 (P200), F123600 (P20), and F144801 (P2)]
Vortex mixer (LMS, model: VTX-3000L)
Insect breeding case (SPL Life Sciences, catalog number: SPL-310102)
Water container (see Figure 2; Umano Kagaku Youki Kabushiki Gaisya, Pla tsubo 20 mL) with a hole (3 mm diameter) on a lid
Figure 2. Post-injection care. (A) Overview of the H. axyridis rearing system. (B) Overview of the aphid rearing. Aphids are reared on the broad bean seedlings in a net cage. (C) Magnified view for aphids.
Paper-based string for water supply (see Figure 2; Morioto Co., Ltd. 2 mm wide) cut at 4 cm long and placed in the water container
Hairdryer (Koizumi, catalog number: KHD-1430 or equivalent)
Clamp for holding hairdryer (Sibata Scientific Technology or equivalent)
Slide glass case (to keep premade egg holder slide clean)
Humid chamber (see Figure 3H)
Procedure
Preparation of an egg holder plate (Figure 3A)
Cut the double-sided tape approximately 1 mm in width and a slightly shorter length than that of a slide glass.
Attach the tape to the edge of the slide glass.
Press down on the tape from above to ensure it adheres evenly.
Peel off the liner from one side.
Store the prepared plate in a slide glass holder to prevent dust accumulation until use.
Figure 3. Injection setup. (A) Alignment of eggs on the double-sided tape of an egg holder plate. (B) Shape of an injection needle. (C and D) Magnified tip of the needle before (C) and after (D) opening. (E) Overview of the microinjection system. The red rectangle highlights the area shown in G. (F) Desiccation system using a hair dryer and adjusting the height with a clamp 20 cm apart from an egg plate positioned underneath. (G) An enlarged view of the microscope stage shows the adjustment of the egg plate height to the injection needle with a glass plate. (H) Overview of the humid chamber. Place a moist paper towel inside. Scale bars in B–D: 100 µm.
Preparation for breeding H. axyridis
Prepare approximately 10 pairs of young ladybird beetles and separate them by one pair per case.
Note: Do not place several pairs in the same case; otherwise, they will compete for mating and will not produce many eggs.
Feed the beetles with plenty of available aphids; they will begin laying eggs after a week of adult emergence. Replace the breeding cases every two days.
Note: While the ladybird beetle can be fed on an artificial diet, a daily supply of fresh aphids is essential for the regular production of eggs.
Microinjection is carried out using eggs from pairs that have been confirmed to lay fertilized eggs.
Preparation of microinjection needle
Injection needles are made by pulling the glass capillary with the following settings (ramp value: 713). This two-step pulling allows the needle to have a longer taper region (Figure 3B).
HEAT 740, PULL 40, VEL 50, TIME 220;
HEAT 740, PULL 40, VEL 60, TIME 220
Store the prepared needles on a needle holder until use.
Egg collection and alignment on the egg holder plate (Figure 3A, Video 1)
On the morning of the day of injection, clean the rearing cases and check for egg laying every 3 h. Microinjection should be done within 3 h after egg laying.
Note: Based on our experience, ladybird beetles like to lay batches of eggs on the back side of the folded pieces of paper or a pea sprig. Eggs laid in these places are easy to collect and handle. The ladybird beetle lays its eggs with their anterior end oriented upward against the surface (paper in the rearing case and leaves in nature), securing them in place using a natural adhesive secreted by the mother onto the posterior end of each egg during oviposition. When enough feeding with aphids is provided, beetles should lay approximately 20–40 eggs per day, even though the timing and location of egg laying are not predictable.
Gently pick up the eggs one by one by softening the egg glue with a moistened paintbrush. Align the eggs on the double-sided tape with the glue side facing outward and spacing them one egg width from each other (Video 1).
Note: The posterior end of the egg is likely to be tougher than the rest of the egg surface, and the egg glue helps to seal the hole made by the injection, contributing to a high survival rate. Furthermore, previous genome editing research using TALEN in Harmonia axyridis has shown that the hatching rate varies significantly depending on the injection site. When injected anteriorly, the hatching rate is 38% (55 out of 144), whereas posterior injection results in a hatching rate of 77% (161 out of 209) [18].
Video 1. Preparation of the egg on a glass slide
Desiccate the eggs with a hair dryer placed 20 cm above in a cool setting for 5 min (Figure 3F).
Preparation of the injection solution and loading it into the injection needle
Centrifuge an injection solution at maximum speed at 4 °C for 3 min. Phosphate-buttered saline or pure water is sufficient for the injection buffer. We recommend avoiding buffers that contain ethylenediaminetetraacetic acid (EDTA), such as Tris-EDTA buffer.
Note: This step is critical for avoiding clogging needle tips during injection.
Carefully load 1.0–1.5 µL of the injection solution from the opposite side of the injection needle with the microloader; 1 µL of injection solution is sufficient to inject approximately 100 eggs.
Note: Make sure the needle does not contain air bubbles. If present, air bubbles could be removed by gently flicking the injection needle.
Set the injection needle to the injection holder and attach the holder to the micromanipulator (Figure 3E), taking care not to damage the needle tip.
Egg microinjection
Set the injection parameter of FemtoJet to a constant pressure (Pc) of 600–700 hPa and an injection pressure of 0 hPa.
Note: In our injection system, the injection solution is constantly leaking from the tip during injection. This is beneficial for preventing the clogging of the needle tips by backflow from the egg yolk or surrounding substrate.
Place the slide glass with aligned eggs onto the stage of the microscope.
The position and height of the needle holder should be adjusted so that the glass needle is almost parallel to the egg holder plate.
Note: Parallel injection is preferred because it ensures that both the needle tip and the egg are focused properly and that an accurate injection is made in the most appropriate location.
Adjust the field of view so that the edge of the egg slide and the needle tip are in the same field of view.
Open the needle tip by gently touching the edge of the slide glass under the microscope; the injection solution will leak out. This process will break the fine end of the needle to be approximately 5–7 µm in diameter (Figure 3C and D, Video 2).
Notes:
After opening the tip, Pc should be adjusted so that a droplet of the leaked injection solution forms at the tip of the needle (Video 3). If the tip is too large, a droplet will not form. This step is critical since injection volume cannot be precisely controlled with this method. We have not measured the maximum volume tolerated by the eggs. It is advisable to first test the needle tip size and duration for which the needle is injected into the eggs to determine the optimal injection conditions.
If the broken tip is too large, it is better to prepare a new needle. Otherwise, such needles will cause significant damage to the egg, leading to reduced survival rates.
Video 2. Preparation of the needle, opening the tip by gently touching the side wall of the glass
Video 3. Microinjecting the solution into the egg of Harmonia axyridis
Under the microscope, move the needle to the center of the field of view and focus on the needle tip; 3–4 eggs fill the field of view.
Move the stage so that the needle is facing the first egg to be injected. It is recommended that injections begin at the end side of the aligned eggs.
Bring the tip of the needle close to the egg surface.
Attach the needle tip to the egg glue, which is localized at the posterior end of the egg, and soften the egg glue by the injection solution leaking out from the tip, allowing the needle tip to penetrate through both the glue and egg membrane into the cytoplasm (Video 3).
Notes:
Once the needle is inserted into the cytoplasm, the injected solution will automatically flow into the cytoplasm. In our experimental setting, we leave the needle for a moment to ensure the solution is flown out into the egg (Video 3). Since the microinjection system cannot precisely control the injection volume, it is important to determine the optimal timing for inserting the needle. Inserting the needle for too long can result in developmental failure.
To provide a better understanding of the approximate injection volume and flow rate, we have recorded a video of the microinjection procedure using a colored injection solution (Video 3). This video allows for a more visual representation of the microinjection process, demonstrating the flow rate and technique more clearly. For first-time microinjections, to better visualize injected eggs, monitor injection and leakage volumes (as shown in Video 3), and ensure accurate injections, we recommend mixing a dye tracer—such as food dye, fluorescent rhodamine, or phenol red—into the injection solution.
Eject the injection needle from the egg and leave the tip of the needle for a moment in the droplet of injection solution outside of the eggshell to prevent the clogging of the solidified yolk.
Continue to use the same needle and repeat the injection with as many eggs as possible. Usually, one needle is used for several glass slides. To obtain the transformant, 100–150 eggs per construct would be enough with appropriate injection skills.
Note: If the needle clogs during injection, pull the needle from the egg and press the CLEAR function or the injection button on the FemtoJet to remove the debris. When the needle tip breaks, enlarging the opening, readjust the Pc value to accommodate the change and maintain proper flow. If this does not work, you may further break the tip or replace it with a new one.
After injection, remove the eggs that were not injected or not properly injected (yolk leakage).
Store the injected eggs with a slide glass in the humid chamber (Figure 3H); then, store the chamber in the incubator at 25 °C just before they hatch.
Note: When we inject water/PBS into the egg using this method, the hatching rate of larvae depends on the injection proficiency, but embryo survival rates typically range from 50% to 80%. Of the hatched larvae, usually approximately 70% develop into adults.
Post-injection care
Just before hatching (approximately 3 days after injection), transfer the glass slide with the injected embryos to an insect case (Figure 2A) and put some frozen H. axyridis eggs as food.
Note: In the insect container, we put a water-absorbing sponge for first- and second-instar larvae and a water container for the remaining stages. For the water container, we use a drill to make a hole in the cap and fill the container with water, inserting twisted paper into the hole.
Feed enough frozen H. axyridis eggs to the hatchling larvae until they reach the second instar, which usually takes three days after hatching (Figure 1).
Note: H. axyridis larvae have an intense cannibalistic habit, so it is important to provide sufficient food after hatching to prevent the loss of injected larvae.
Once larvae reach the third instar (Figure 1; 6–7 days after hatching), separate them into five larvae per case and feed them with frozen or fresh aphids daily (Figure 2).
From the fourth instar onward, the larvae can be reared on artificial food.
Note: Once the larvae have reached the fourth-instar stage, they should be separated into one larva per case to prevent them from cannibalizing each other.
Validation of protocol
This protocol or parts of it has been used and validated in the following research articles:
Kuwayama et al. [15]. Germ‐line transformation and RNAi of the ladybird beetle, Harmonia axyridis. Insect Mol Biol. (Figure 2)
Kuwayama et al. [14]. Establishment of Transgenic Lines for Jumpstarter Method Using a Composite Transposon Vector in the Ladybird Beetle, Harmonia axyridis. PLoS One. (Figure 1)
Osanai-Futahashi et al. [16]. A visible dominant marker for insect transgenesis. Nat Commun. (Figure 6c)
Hatakeyama et al. [18]. Knockout of a transgene by transcription activator‐like effector nucleases (TALENs) in the sawfly, Athalia rosae (Hymenoptera) and the ladybird beetle, Harmonia axyridis (Coleoptera). Insect Mol Biol. (Figure 3B)
Acknowledgments
We extend our gratitude to the Emerging Model Organisms Facility of NIBB Trans-Scale Biology Center for technical assistance. This protocol was adapted from Kuwayama et al. [14,15], Osanai-Futahashi et al. [16], and Hatakeyama et al. [18]. This work was supported by JSPS KAKENHI Grant Numbers 21H02211 to T. Nakamura, 22K15131 to Y. Matsuoka, and 16H02596, 19H01004, and 23H02227 to T. Niimi.
Competing interests
No conflict of interest is declared with regard to this article.
Ethical considerations
No ethical requirements are needed with regard to this article.
References
Komai, T. (1956). Genetics of Ladybeetles. In: Demerec, M. (Ed.). Advances in Genetics (Vol. 8, pp. 155–188). Academic Press.
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Publication history
Received: May 24, 2024
Accepted: Oct 10, 2024
Available online: Oct 30, 2024
Published: Dec 20, 2024
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© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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5,137 | https://bio-protocol.org/en/bpdetail?id=5137&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
A Bio-protocol resource
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Muscle Biopsy Sample Preparation and Proteomics Analysis Based on UHPLC-MS/MS
JD Jiawei Du *
JH Jinghua Hou *
HY Hezhang Yun *
YS Yafeng Song
(*contributed equally to this work)
Published: Vol 14, Iss 24, Dec 20, 2024
DOI: 10.21769/BioProtoc.5137 Views: 240
Reviewed by: Amit Kumar Dey Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Molecular & Cellular Proteomics Apr 2024
Abstract
Proteomics analysis is crucial for understanding the molecular mechanisms underlying muscle adaptations to different types of exercise, such as concentric and eccentric training. Traditional methods like two-dimensional gel electrophoresis and standard mass spectrometry have been used to analyze muscle protein content and modifications. This protocol details the preparation of muscle samples for proteomics analysis using ultra-high-performance liquid chromatography (UHPLC). It includes steps for muscle biopsy collection, protein extraction, digestion, and UHPLC-based analysis. The UHPLC method offers high-resolution separation of complex protein mixtures, providing more detailed and accurate proteomic profiles compared to conventional techniques. This protocol significantly enhances sensitivity, reproducibility, and efficiency, making it ideal for comprehensive muscle proteomics studies.
Key features
• Developed for analyzing muscle adaptations in response to concentric and eccentric training, applicable to various physiology exercise studies.
• Utilizes UHPLC-MS/MS for high-resolution separation and detailed proteomic profiling.
• Requires access to advanced UHPLC-MS/MS equipment and muscle biopsy collection tools.
• The protocol can be completed within one week, including sample preparation and analysis.
Keywords: Muscle proteomics UHPLC-MS/MS Concentric exercise Eccentric exercise Muscle biopsy
Graphical overview
Background
Proteomics is an essential field in biomedical research, offering deep insights into the molecular mechanisms underlying various physiological and pathological processes [1]. This protocol focuses on muscle proteomics, specifically analyzing muscle adaptations to different types of exercise, such as concentric (CON) and eccentric (ECC) training. Muscle proteomics can elucidate the intricate molecular changes that occur in response to physical activity, thereby advancing our understanding of exercise physiology, muscle hypertrophy, and recovery [2]. Traditional liquid chromatography–mass spectrometry (LC–MS) techniques have long been employed for proteomic analysis, offering significant advantages over two-dimensional gel electrophoresis in terms of sensitivity and reproducibility [3]. However, the separation power of conventional LC systems often falls short when dealing with complex mixtures like muscle tissue proteomes, where resolving peptides with close retention times and molecular weights is crucial [4].
The protocol described here utilizes ultra-high-performance liquid chromatography (UHPLC) for the preparation and analysis of muscle samples. This method offers several significant advantages over traditional approaches. UHPLC provides superior resolution and separation of complex protein mixtures compared to conventional liquid chromatography, allowing for more detailed and accurate proteomic profiling [5]. The protocol significantly increases the sensitivity of protein detection, enabling the identification of low-abundance proteins that might be missed using standard methodologies [6]. Additionally, the UHPLC method enhances reproducibility in proteomic analysis due to its precise control over chromatographic conditions and consistent performance across different runs [7]. This protocol also reduces the time required for sample preparation and analysis compared to traditional methods, allowing for quicker turnaround and higher throughput.
Despite its advantages, the UHPLC-based protocol has some limitations. The protocol requires access to advanced UHPLC equipment, which may not be available in all research laboratories. Implementing this protocol necessitates a certain level of technical expertise in UHPLC and proteomic analysis, which may require specialized training for some laboratory personnel. Beyond muscle proteomics, this protocol can be adapted for use in other research areas that involve complex protein mixtures. For example, researchers can apply this protocol to nutritional studies, examining proteomic changes in muscle tissues in response to different dietary interventions and providing insights into the effects of nutrition on muscle health and function [8]. Additionally, the protocol can be used to evaluate the molecular effects of pharmacological agents on muscle tissues, aiding in the development of new drugs aimed at treating muscle-related diseases or enhancing muscle performance [9].
In summary, the UHPLC-based protocol for muscle proteomics offers significant improvements in resolution, sensitivity, reproducibility, and efficiency over traditional methodologies. While it does require specialized equipment and expertise, its advantages make it a powerful tool for advancing research in exercise physiology, muscle adaptation, and a wide range of other biomedical fields. This protocol not only enhances our understanding of molecular adaptations to exercise but also holds promise for broader applications in disease research, nutritional science, and drug development.
Materials and reagents
Biological materials
Human vastus lateralis muscle biopsy sample
Reagents
Povidone-iodine (Sigma-Aldrich, catalog number: PVP-I)
2% lidocaine with epinephrine (Hospira, catalog number: 0409-4279-16)
Glucose solution (Sigma-Aldrich, catalog number: G8769)
Physiological saline (Sigma-Aldrich, catalog number: S8776)
Liquid nitrogen (Air Products, catalog number: NI)
Isopentane (Sigma-Aldrich, catalog number: 277258)
Tris-HCl (Sigma-Aldrich, catalog number: T5941)
Triton X-100 (Sigma-Aldrich, catalog number: T8787)
Protease inhibitor cocktail (Sigma-Aldrich, catalog number: P2714)
BCA Protein Assay Kit (Thermo Fisher, catalog number: 23227)
Trichloroacetic acid (TCA) (Sigma-Aldrich, catalog number: T6399)
Acetone, HPLC grade (Sigma-Aldrich, catalog number: 34850)
Triethylammonium bicarbonate (TEAB) (Sigma-Aldrich, catalog number: T7408)
Trypsin (Promega, catalog number: V5111)
Dithiothreitol (DTT) (Sigma-Aldrich, catalog number: D0632)
Iodoacetamide (IAA) (Sigma-Aldrich, catalog number: I1149)
Formic acid (FA), HPLC grade (Sigma-Aldrich, catalog number: 56302)
Acetonitrile (ACN), HPLC grade (Sigma-Aldrich, catalog number: 34967)
Water, HPLC grade (Thermo Fisher, catalog number: W7-4)
EASY-nLC 1200 solvent A (0.1% formic acid in water) (Thermo Fisher, catalog number: LS1224)
EASY-nLC 1200 solvent B (0.1% formic acid in acetonitrile) (Thermo Fisher, catalog number: LS1225)
FAIMS Pro compensation voltage solution (Thermo Fisher, catalog number: FMSP 0003)
Solutions
PBS (see Recipes)
Triton lysis buffer (see Recipes)
LC-MS/MS mobile phase A (see Recipes)
LC-MS/MS mobile phase B (see Recipes)
Recipes
PBS
Reagent Final concentration Quantity or Volume
NaCl 137 mM 8.01 g
KCl 2.7 mM 0.2 g
NaHPO 10 mM 1.42 g
KHPO 1.8 mM 0.24 g
Distilled H2O n/a Up to 1 L
Triton lysis buffer
Reagent Final concentration Quantity or Volume
Tris-HCl 50 mM 60.57 mg
NaCl 150 mM 87.66 mg
EDTA 1 mM 0.292 mg
Triton X-100 1% 10 μL
Protease inhibitor 1% 10 μL
Distilled H2O n/a Up to 1 mL
LC-MS/MS mobile phase A
Reagent Final concentration Quantity or Volume
Formic acid 0.1% 1 mL (for 1 L)
Acetonitrile 2% 20 mL (for 1 L)
Water -- Up to 1 L
LC-MS/MS mobile phase B
Reagent Final concentration Quantity or Volume
Formic acid 0.1% 1 mL (for 1 L)
Acetonitrile 90% 900 mL (for 1 L)
Water -- Up to 1 L
Laboratory supplies
Disposable semi-automatic biopsy needle (Bard, catalog number: 1520-5010)
Pre-cooled tweezers (VWR, catalog number: 82027-530)
Aluminum foil (Reynolds, catalog number: 614)
Liquid nitrogen storage container (Thermo Fisher, catalog number: 11887163)
Cryogenic vials (Nalgene, catalog number: 5000-1020)
Mortar and pestle set (Bel-Art, catalog number: H37260-0000)
Microcentrifuge tubes (Eppendorf, catalog number: 0030120086)
1.5 mL microcentrifuge tubes (Eppendorf, catalog number: 022363204)
50 mL conical tubes (Falcon, catalog number: 352070)
1,000 μL pipette tips (Rainin, catalog number: RT-L1000F)
200 μL pipette tips (Rainin, catalog number: RT-L200F)
10 μL pipette tips (Rainin, catalog number: RT-L10F)
Sterile gloves (Medline, catalog number: MDS195285)
Sterile surgical drapes (3M, catalog number: 1010)
Sterile gauze pads (Dynarex, catalog number: 3344)
Surgical mask (Medline, catalog number: NON27378)
Medical adhesive tape (3M, catalog number: 1530-1)
Medical bandages (Johnson & Johnson, catalog number: 2015)
96-well plates (Thermo Fisher, catalog number: 260836)
Equipment
Ultrasound machine (GE Healthcare, model: LOGIQ E9)
-80 °C freezer (Thermo Fisher, model: ULT2186)
Centrifuge (Eppendorf, model: 5424)
Ultrasonic homogenizer (Qsonica, model: Q55)
Vortex mixer (VWR, model: 945302)
UHPLC system (Thermo Fisher, model: EASY-nLC 1200)
nanoACQUITY HPLC HHS T3 column (Waters, catalog number: 186003538)
Mass spectrometer (Thermo Fisher, model: Orbitrap Exploris 480)
FAIMS Pro Interface (Thermo Fisher, catalog number: FAIMS01-10000)
Water bath (Grant Instruments, model: JB Academy)
pH meter (Mettler Toledo, model: SevenCompact)
Magnetic stirrer (IKA, model: RCT basic)
Analytical balance (Sartorius, model: Entris II)
Refrigerated microcentrifuge (Eppendorf, model: 5418 R)
Electronic pipettes (Eppendorf, model: Xplorer plus)
Software and datasets
UniProt (https://www.uniprot.org/) (Access date, 2021/08)
Proteome Discoverer (v2.4.1.15, Thermo Scientific)
Procedure
Muscle biopsy
Preparation
Ensure all equipment and reagents are prepared and sterile.
Set up a sterile area and prepare a disposable semi-automatic biopsy needle.
Biopsy procedure
Position the subject comfortably and identify the biopsy site (typically the vastus lateralis muscle).
Disinfect the target puncture site three times using povidone-iodine, expanding the circular disinfection area around the puncture point. Inject 3–5 mL of 2% lidocaine containing 50 μL of epinephrine for local anesthesia of the skin and subcutaneous tissue. Ensure the needle does not penetrate the deep fascia to avoid anesthetic contamination of muscle tissue. Provide the subject with sufficient glucose to prevent hypoglycemia and stabilize their emotions.
The physician holds the disposable semi-automatic biopsy needle and slowly inserts the needle through the skin and subcutaneous tissue, using musculoskeletal ultrasound to guide the needle and avoid important nerves and blood vessels. Upon reaching the deep fascia, increase the force and rotate the needle to penetrate the fascia. Use ultrasound to confirm the needle's position in the vastus lateralis muscle. The assistant gently compresses the muscle around the puncture site, while the physician triggers the needle mechanism to cut the muscle (Figure 1A).
Figure 1. Muscle biopsy. A. Biopsy procedure. B. Muscle sample preservation.
Given the small diameter and cutting window of the biopsy needle, multiple cuts (5–7 times) may be necessary to obtain sufficient muscle tissue (~50 mg).
Sample preservation
Remove the muscle tissue from the biopsy needle using pre-cooled tweezers, rinse multiple times with saline, wrap in foil, quickly freeze in liquid nitrogen–cooled isopentane, and temporarily store in a liquid nitrogen container. Finally, store all muscle samples in a -80 °C freezer for proteomic analysis (Figure 1B).
Post-procedure care
Remove the biopsy needle and apply a medical dressing to the puncture site.
Instruct the subject to sit quietly for 15–20 min, applying continuous pressure to the puncture site to prevent bleeding. Advise the subject not to shower for 24 h and to keep the wound clean and dry.
Continuously monitor and record the subject's condition and wound recovery until fully healed.
Preparation of muscle samples for proteomic analysis
Homogenization
Retrieve muscle samples from the -80 °C freezer and place them on ice.
Weigh the tissue sample into a liquid nitrogen–cooled mortar and grind it into a fine powder.
Lysis
Add four times the volume of lysis buffer (see Recipes) to the powdered muscle samples. Perform ultrasonic lysis to homogenize the muscle tissue.
Centrifuge at 12,000× g for 10 min at 4 °C, discard the pellet, and transfer the supernatant to a new tube.
Determine protein concentration using a BCA assay kit.
Protein digestion
Digest equal amounts of muscle sample protein from each group (100 μg to 1 mg). Adjust the volume with lysis buffer.
Slowly add TCA to a final concentration of 20%, vortex to mix, and precipitate at 4 °C for 2 h.
Centrifuge at 4,500× g for 5 min at 4 °C, discard the supernatant, and wash the pellet 2–3 times with pre-cooled acetone.
Air-dry the pellet (15–30 min), add TEAB to a final concentration of 200mM, and sonicate to disperse the pellet. Sonication time is 10–30 s per cycle; power is 30%–50% output; 3–5 cycles with 30 s to 1 min cooling on ice between cycles.
Add DTT to a final concentration of 5 mM and reduce at 56 °C for 30 min.
Add IAA to a final concentration of 11 mM and incubate in the dark at room temperature for 15 min.
Add trypsin at a 1:50 ratio (enzyme:protein, m/m) and digest overnight at 37 °C.
UHPLC-MS/MS analysis
Calibration process of UHPLC-MS/MS
Calibration of the instrument: To ensure accuracy, the UHPLC-MS/MS system is typically calibrated by injecting standard solutions with known concentrations. The calibration process aims to validate the performance of chromatographic separation and mass spectrometric detection and ensure a good linear response across the concentration range of the target analyte.
Optimization of mass spectrometry parameters: Calibration of the mass spectrometry component involves optimizing ion source conditions (e.g., spray voltage, ion source temperature) and settings of the mass analyzer to ensure accurate detection of compounds across different molecular weights, with stable peak shapes and responses.
Mass calibration: Using standard mass calibration substances (such as known peptides or small molecules), the mass accuracy of the spectrometer is checked regularly, and necessary corrections are made to ensure that the mass deviation is within the permissible range.
Preparation of calibration solutions
Selection of standards: The standards used in the experiment should be high-purity target compounds, usually with a purity of over 98%. These standards could be purified forms of the target analyte, isotope-labeled compounds, or peptides with known sequences.
Preparation of stock solutions: First, weigh the standard and dissolve in an appropriate solvent (e.g., 0.1% formic acid solution or acetonitrile) to prepare a high-concentration stock solution. The stock solution should have an accurate concentration, typically 1 mg/mL or higher.
Dilution of standard solutions: From the stock solution, prepare a series of diluted solutions of different concentrations to cover the expected concentration range of the samples. The concentration of the standard solutions usually ranges from 1 ng/μL to 500 ng/μL, depending on the experimental requirements.
Establishment of calibration standards
Calibration curve: Inject different concentrations of the standard solutions into the UHPLC-MS/MS system, record the mass spectrometric response (such as peak area or peak height) for each concentration, and plot a calibration curve. The calibration curve should exhibit good linearity across the measured concentration range to ensure the accuracy of quantitative analysis.
Use of internal standards: To compensate for variability during sample preparation and detection, internal standards (such as isotope-labeled peptides or compounds) may be used. Internal standards can be added to both the sample and the standard solution to correct for experimental errors.
Quality control and validation
Regular calibration: The calibration process should be conducted regularly, especially if the instrument undergoes maintenance or there are changes in experimental conditions. Regular validation of the instrument’s performance ensures consistency and reliability in calibration results.
Use of quality control samples: Quality control (QC) samples can be used to assess the instrument’s performance and ensure that the mass spectrometric response falls within the expected range.
Validation of calibration results
Linear validation: Ensure that the calibration curve maintains a good linear relationship across the entire analytical range (usually requiring an R2 greater than 0.99).
Limit of detection (LOD) and limit of quantification (LOQ): Based on the calibration curve, determine the instrument’s LOD and LOQ to ensure that accurate detection results can still be obtained at low concentrations.
Sample loading and separation
Inject 1 μg of peptide dissolved in 0.1% formic acid into an EASY-nLC1200 system using a nanoACQUITY HPLC HHS T3 column (75 μm × 150 mm, 1.8 μm). The mobile phase A is 0.1% formic acid with 2% acetonitrile; mobile phase B is 0.1% formic acid with 90% acetonitrile.
Set the gradient: 0–68 min, 6%–23% B; 68–82 min, 23%–32% B; 82–86 min, 32%–80% B; 86–90 min, 80% B. Maintain the flow rate at 450 nL/min.
Ionization and mass spectrometry
The separated peptides are ionized using an NSI source and analyzed by an Orbitrap Exploris 480 mass spectrometer. Set the capillary temperature to 260 °C and ionization voltage to 2.3 kV. The FAIMS compensation voltage (CV) is set to -45V, -65V.
Mass spectrometry analysis
Both the parent ions and their fragments are detected using the high-resolution Orbitrap. The primary scan range is 400–1,200 m/z with a resolution of 60,000; the secondary scan range starts at 110 m/z with a resolution of 15,000. TurboTMT is off.
Data acquisition
Use data-dependent acquisition (DDA) to select the top 25 precursor ions for fragmentation using HCD with a collision energy of 27% and secondary analysis.
To optimize the use of the mass spectrometer, set the automatic gain control (AGC) to 100%, the signal threshold to 5E4 ions/s, maximum injection time to Auto, and dynamic exclusion to 20 s.
Data analysis
Mass spectrometry data analysis
Quality control
Peptide length distribution: Most effective peptides should be between 7 and 20 amino acids in length, with the ideal length being 8–15 amino acids (Figure 2A).
Figure 2. Quality control. A. Peptide length distribution. The colors represent the amount of charge carried by the peptide. B. Peptide count distribution. C. Protein coverage distribution. The colors represent protein coverage. D. Protein molecular weight distribution.
Peptide count distribution: At least 2–3 unique peptides should be detected for each protein to ensure reliability and reproducibility (Figure 2B).
Protein coverage distribution: Generally, the protein coverage should exceed 20%–30%, with high-abundance proteins ideally achieving 50% or more (Figure 2C).
Protein molecular weight distribution. A wide range of molecular weights should be covered, typically from 5 to 250 kDa, ensuring the detection of proteins of various sizes within this range (Figure 2D).
Filter data at the spectrum, peptide, and protein levels with an FDR of 1%.
Data processing
Use Proteome Discoverer (v2.4.1.15) to analyze the secondary mass spectrometry data. Set the database to Homo_sapiens_9606_SP_20210721.fasta (20,387 sequences) with a decoy database to calculate the false discovery rate (FDR) and a contaminant database to eliminate common contaminants. Set enzyme to Trypsin (Full) with up to two missed cleavages, minimum peptide length of six residues, and a maximum of three modifications per peptide. Set the precursor mass tolerance to 10 ppm and fragment mass tolerance to 0.02 Da.
The mass tolerance for precursor ions is set to 10 ppm, and to 0.02 Da for fragment ions; the maximum allowed retention time deviation is 10 min.
Search engines choose Sequest HT. Set Carbamidomethyl (C) as a fixed modification; Oxidation (M), Acetyl (N-terminus), Met-loss (M), Deamidated (N, Q), and Met-loss + acetyl (M) as variable modifications. Set FDR for protein, peptide, and PSM identification to 1%.
In the Minora detection process, set the minimum trace length for signal identification to 5, Max ΔRT of Isotope Pattern Multiplets [min] to 0.02, and the confidence of the Peptide-Spectrum Match to HIGH.
Use UNIQUE and RAZOR peptides for quantification based on intensity values. Protein abundance was calculated as summed abundance.
Quantitative analysis
Normalization
Normalize raw mass spectrometry data both row-wise and column-wise. Divide each protein's quantification value by its average value across all samples (row-wise). Then, divide each protein in a sample by the median value of that sample (column-wise) to obtain normalized data for bioinformatics analysis.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Du et al. [10]. Proteomic Profiling of Muscular Adaptations to Short-Term Concentric Versus Eccentric Exercise Training in Humans. Mol Cell Proteomics (Supplemental Table S2)]
General notes and troubleshooting
General notes
Problem 1: How to recalibrate the gradient and how to optimize the column temperature.
Recalibrate the gradient
Check the gradient program: Begin by reviewing the existing gradient program to determine whether it is appropriate for separating the target compounds. If the gradient time or solvent ratio is incorrect, it may lead to retention time shifts or poor separation of the target analytes.
Optimize start and end conditions: Adjust the starting and ending solvent ratios based on the polarity and chemical nature of the sample. For polar compounds, use a higher proportion of the aqueous phase at the start. For hydrophobic compounds, start with more of the organic phase.
Extend or shorten gradient time: If peaks are not well-separated, consider extending the gradient time to allow more time for different compounds to separate. If the peaks are too broad, resulting in lower resolution, shorten the gradient time.
Validate with standards: Use known standard samples to check if the new gradient can effectively separate the target compounds and ensure stable retention times. After adjusting the gradient, record the retention time for each compound and ensure it remains consistent.
Optimize column temperature
Determine the optimal temperature range: Typically, column temperature is optimized between 25 °C and 60 °C. Higher temperatures can speed up the analysis process and reduce retention times but may lead to decreased resolution. Lower temperatures, on the other hand, increase resolution but result in longer analysis times.
Monitor peak shape and resolution: Gradually adjust the temperature (e.g., by increasing or decreasing it by 5 °C at a time) and observe changes in peak shape and resolution. Generally, as temperature increases, sample mobility improves, and peaks become narrower. However, excessive temperature increases may lead to peak fronting or peak loss.
Balance analysis time and resolution: Select a temperature that maintains good separation while also not excessively prolonging the analysis time, ensuring sufficient resolution.
Consider sample stability: For heat-sensitive samples, avoid using excessively high column temperatures to prevent sample degradation in the column.
Problem 2: How to increase the sample volume within the linear range of the instrument.
In LC–MS methods, increasing the sample amount must ensure it remains within the linear range of the instrument to avoid signal saturation and ensure quantitative accuracy. The following steps and considerations should be followed when increasing the sample amount:
Confirm the instrument's linear range: The linear range of the instrument refers to the range in which the analyte concentration has a linear relationship with the detection signal (e.g., peak area). In this range, the signal increases linearly with the sample concentration. Create a calibration curve by injecting standards at different concentrations to determine the linear range. This curve should cover the expected sample concentrations and typically uses at least five different concentration points.
Gradually increase sample amount: If the sample amount needs to be increased, it is recommended to gradually increase the concentration or injection volume while monitoring the signal response. Stop increasing the sample amount before the signal enters the nonlinear range. The sample amount can be increased by injecting a larger volume into the liquid chromatography system, for example, increasing from 10 to 20 μL. However, ensure that the injection volume does not exceed the sample loop capacity.
Evaluate signal saturation: While increasing the sample amount, monitor the signal intensity of the mass spectrometer. If the signal intensity starts to show nonlinear growth or approaches the saturation point, reduce the sample concentration or injection volume accordingly. Increasing the sample amount may lead to increased co-eluting substances, causing ion suppression and affecting the detection of analytes. Therefore, closely monitor for ion suppression and adjust sample preparation methods if necessary.
Appropriate sample dilution: For high-concentration samples, perform dilution to ensure that the sample concentration falls within the instrument's linear range. The dilution factor should be chosen based on the linear range of the calibration curve. Internal standards can help correct signal deviations that may result from increased sample amounts, ensuring the accuracy of the quantification results.
Optimize mass spectrometer parameters: When increasing the sample amount, adjust the ion source parameters of the mass spectrometer (e.g., spray voltage, gas flow, temperature) to prevent ion source overload. By optimizing the scan range or using selected reaction monitoring (SRM) mode, focus on the target analytes and minimize interference from other substances.
Maintain good peak shape: Increasing the sample amount may cause peak broadening, affecting the separation. Ensure that the column selection is appropriate, and that flow rate, gradient, and other parameters are optimized to maintain good peak shape. Excessive sample amounts may overload the column, leading to peak broadening or tailing. Monitor the peak shape when increasing the sample amount to ensure the column is not overloaded.
Repeat experiments and validation: After each increase in sample amount, repeat the experiment to ensure consistency and reproducibility, avoiding random deviations. After increasing the sample amount, validate the calibration curve again to confirm that the linear range and detection limits remain unaffected.
Troubleshooting
Problem 1: Poor chromatographic separation.
Possible cause: Incorrect solvent composition or gradient program.
Solution: Double-check the solvent preparation and gradient program. Ensure that solvent bottles are correctly connected and that the mobile phase composition matches the method. If the problem persists, consider recalibrating the gradient and optimizing the column temperature.
Problem 2: Low signal intensity in MS.
Possible cause: Insufficient sample loading.
Solution: Increase the sample injection volume within the linear range of the instrument. Ensure that samples are properly dissolved in the appropriate solvent. Check for clogs or blockages in the sample introduction system and clean or replace them as necessary.
Problem 3: Poor ionization efficiency.
Possible cause: Contaminated ion source or electrospray emitter.
Solution: Clean the ion source and electrospray emitter according to the manufacturer's instructions. Use appropriate solvents for cleaning to avoid damaging sensitive components. Regularly perform maintenance procedures recommended by the instrument manual to ensure optimal ionization conditions.
Acknowledgments
This study was supported by the National Natural Science Foundation of China (82071413, 82271438). This protocol was adapted and modified from Du et al. [10].
Competing interests
The authors declare no competing interests.
Ethical considerations
This protocol was conducted according to the Declaration of Helsinki's ethical principles. The local ethics committee of Sports Science Experimental Ethics Committee at Beijing Sport University approved the study (Permission number: 2020148H).
References
Anderson, N. L. and Anderson, N. G. (2002). The Human Plasma Proteome. Mol Cell Proteomics. 1(11): 845–867.
Hoffman, N. J., Parker, B. L., Chaudhuri, R., Fisher-Wellman, K. H., Kleinert, M., Humphrey, S. J., Yang, P., Holliday, M., Trefely, S. and Fazakerley, D. J. (2015). Global Phosphoproteomic Analysis of Human Skeletal Muscle Reveals a Network of Exercise-Regulated Kinases and AMPK Substrates. Cell Metab. 22(5): 922–935.
Görg, A., Weiss, W. and Dunn, M. J. (2004). Current two‐dimensional electrophoresis technology for proteomics. Proteomics. 4(12): 3665–3685.
Aebersold, R. and Mann, M. (2003). Mass spectrometry-based proteomics. Nature. 422(6928): 198–207.
Howard, J. W., Kay, R. G., Pleasance, S. and Creaser, C. S. (2012). UHPLC for the separation of proteins and peptides. Bioanalysis. 4(24): 2971–2988.
Smaczniak, C., Li, N., Boeren, S., America, T., van Dongen, W., Goerdayal, S. S., de Vries, S., Angenent, G. C. and Kaufmann, K. (2012). Proteomics-based identification of low-abundance signaling and regulatory protein complexes in native plant tissues. Nat Protoc. 7(12): 2144–2158.
Perez de Souza, L., Alseekh, S., Scossa, F. and Fernie, A. R. (2021). Ultra-high-performance liquid chromatography high-resolution mass spectrometry variants for metabolomics research. Nat Methods. 18(7): 733–746.
Tipton, K. D., Elliott, T. A., Cree, M. G., Aarsland, A. A., Sanford, A. P. and Wolfe, R. R. (2007). Stimulation of net muscle protein synthesis by whey protein ingestion before and after exercise. Am J Physiol Endocrinol Metab. 292(1): E71–E76.
Yarasheski, K. E., (2003). Review Article: Exercise, Aging, and Muscle Protein Metabolism. J Gerontol A Biol Sci Med Sci. 58(10): M918–M922.
Du, J., Yun, H., Wang, H., Bai, X., Su, Y., Ge, X., Wang, Y., Gu, B., Zhao, L. and Yu, J. G. (2024). Proteomic Profiling of Muscular Adaptations to Short-Term Concentric Versus Eccentric Exercise Training in Humans. Mol Cell Proteomics. 23(4): 100748.
Article Information
Publication history
Received: Jun 19, 2024
Accepted: Oct 10, 2024
Available online: Oct 31, 2024
Published: Dec 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Cell Biology > Cell-based analysis > Protein secretion
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An HPLC-based Assay to Study the Activity of Cyclic Diadenosine Monophosphate (C-di-AMP) Synthase DisA from Mycobacterium smegmatis
AM Avisek Mahapa *
SG Sudhanshu Gautam *
AR Arti Rathore
DC Dipankar Chatterji
(*contributed equally to this work)
Published: Vol 14, Iss 24, Dec 20, 2024
DOI: 10.21769/BioProtoc.5138 Views: 277
Reviewed by: Ritu GuptaPrajita PandeySoumya Moonjely
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Original Research Article:
The authors used this protocol in Protein Science Jan 2023
Abstract
Cyclic diadenosine monophosphate (c-di-AMP) is a recently discovered second messenger that modulates several signal transduction pathways in bacterial and host cells. Besides the bacterial system, c-di-AMP signaling is also connected with the host cytoplasmic surveillance pathways (CSP) that induce type-I IFN responses through STING-mediated pathways. Additionally, c-di-AMP demonstrates potent adjuvant properties, particularly when administered alongside the Bacillus Calmette–Guérin (BCG) vaccine through mucosal routes. Because of its pivotal role in bacterial signaling and host immune response, this molecule has garnered significant interest from the pharmaceutical industry. This protocol outlines the quantification of c-di-AMP by an HPLC-based assay to enumerate the activity of c-di-AMP synthase from Mycobacterium smegmatis. The following protocol is designed to be generic, enabling the study of c-di-AMP synthase activity from other bacterial species. However, modifications may be required depending on the specific activity of c-di-AMP synthase from different bacterial sources.
Key features
• Easy and time-saving HPLC-based quantification method for c-di-AMP.
• Quick and reliable method to study enzymatic activity/kinetics of DisA/DAC.
• Elimination of potentially hazardous radioactive substrates and products for c-di-AMP quantification.
Keywords: C-di-AMP Secondary messenger Mycobacteria DisA Diadenylate cyclase (DAC) HPLC Quantification
Graphical overview
C-di-AMP homeostasis in mycobacteria and HPLC-based quantification process. A. Overview of c-di-AMP synthesis and hydrolysis in M. smegmatis. B. Overview of DisA protein purification, in vitro C-di-AMP synthesis, and HPLC-based quantification process.
Background
Cyclic di-adenosine monophosphate (c-di-AMP) is an essential secondary messenger that plays a pivotal role in regulating various physiological processes, including osmoregulation, DNA integrity maintenance, sporulation, cell wall homeostasis, ion-channel regulation, antibiotic resistance, virulence gene expression, acid resistance, and carbon metabolism [1,2]. In the bacterial kingdom, c-di-AMP synthesis is catalyzed by di-adenylate cyclase (DAC) domain–containing proteins, which convert two ATP molecules into c-di-AMP in the presence of metal ions such as Mg and Mn. To date, five classes of DAC domain–containing proteins have been identified in bacteria: DNA integrity scanning protein A (DisA), c-di-AMP synthase A (CdaA), c-di-AMP synthase sporulation-specific (CdaS), c-di-AMP synthase N-terminal TM segment (CdaM), and c-di-AMP synthase Z (CdaZ) [3,4]. DisA, an octameric protein, has dual functions: it can bind to Holliday junction DNA or synthesize c-di-AMP [1]. Mycobacterium smegmatis is a non-pathogenic, rapidly growing bacterium from the genus Mycobacterium. It serves as a widely used model organism for studying mycobacteria, including pathogenic species like Mycobacterium tuberculosis and non-tuberculous mycobacteria (NTM) pathogens, such as Mycobacterium abscessus and Mycobacterium avium [5]. In M. smegmatis, the gene MSMEG_6080 encodes the c-di-AMP synthase DisA, which catalyzes c-di-AMP synthesis in the presence of Mg ions. DisA contains three main domains: the N-terminal DAC domain-1, the C-terminal DNA binding domain-3, and a linker domain (domain-2) that connects domain 1 and domain 2 [4,6]. The enzyme's active site is located at the interface of the DAC domains [1]. The cellular concentration of c-di-AMP is regulated by phosphodiesterases (PDEs), which are located in different operons and contain DHH-DHHA1 or HD (His-Asp) domains [7–9]. PDEs hydrolyze c-di-AMP into pApA or AMP. These enzymes typically feature a unique structural alignment, consisting of an N-terminal linked to a degenerate GGDEF domain and a C-terminal DHH-DHHA1 module, essential for c-di-AMP hydrolysis [9]. In Mycobacterium tuberculosis, the PDE (CnpB) has a core DHH-DHHA1 domain that hydrolyzes c-di-AMP first to linear 5'-pApA and then to two 5'-AMP molecules [8]. Similarly, the PDE in M. smegmatis (MSMEG_2630) contains a DHH-DHHA1 domain without additional regulatory domains found in the GdpP protein family of Bacillus subtilis. All types of PDEs require specific divalent metal ions (Mg, Mn, Co) for their activity [1,2,7]. The opposing activities of these enzymes maintain the homeostasis of c-di-AMP within bacterial cells. Herein, we describe an HPLC-based protocol to study the c-di-AMP synthesis by DisA protein from M. smegmatis. This protocol outlines a straightforward method for quantifying c-di-AMP synthesized by mycobacterial DisA. The method is adapted from protocols by Bai et al., Christen et al., and Ryjenkov et al., with necessary modifications [10–13]. This HPLC-based method allows quick, easy, and quantitative estimation of synthesized c-di-AMP by DisA, avoiding the utilization of potentially hazardous radioactive substrates and products.
Materials and reagents
Reagents
Purified DisA protein (Origin: M. smegmatis)
ATP (Sigma, catalog number: A2383)
C-di-AMP (Jena Bioscience, catalog number: NU-954S)
Tris base (Sigma, catalog number: 10708976001)
EDTA (Sigma, catalog number: E4884)
Sodium chloride (NaCl) (Sigma, catalog number: S9888)
Tetrabutylammonium hydrogen sulfate (Sigma, catalog number: 15583)
Methanol (HPLC grade) (Sigma, catalog number: 34860)
Double-distilled water (Milli-Q Ultrapure Water Systems)
KH2PO4 (Sigma, catalog number: P5655)
MgCl2 (Sigma, catalog number: M8266)
Tris-Cl (Sigma, catalog number: 10812846001)
Luria Bertani Broth, Miller (HIMEDIA, catalog number: M1245)
IPTG (Sigma, catalog number I6758)
Laboratory supplies
Dialysis membrane (Sigma, catalog number: D9527)
Pipette tips (Tarsons, catalog numbers: 521000, 521010, 521020)
1.5 mL reaction tubes (Tarsons, catalog number: 500010)
Membrane filter, 0.22 μm pore size (Merck, catalog number: GSWP04700)
Solutions
Protein purification buffers: lysis buffer, wash buffer, and elution buffer (see Recipes)
DisA dialysis buffer (buffer A) (see Recipes)
Size exclusion chromatography (SEC) (buffer B) (see Recipes)
Buffer C for c-di-AMP synthesis reaction (see Recipes)
Buffer D (see Recipes)
Buffer E (see Recipes)
Recipes
Protein Purification buffers
Lysis buffer
50 mM Tris-Cl (pH 7.9), 300 mM NaCl, and 1 mM phenylmethylsulfonyl fluoride (PMSF)
Wash buffer
50 mM Tris-Cl (pH 7.9), 300 mM NaCl, and 40 mM imidazole
Elution buffer
50 mM Tris-Cl (pH 7.9), 300 mM NaCl, and 300 mM imidazole
DisA dialysis buffer (buffer A)
50 mM Tris-Cl buffer at pH 7.5, 300 mM NaCl
Size exclusion chromatography (SEC) (buffer B)
50 mM Tris-Cl (pH 7.9), 300 mM NaCl
Buffer C for c-di-AMP synthesis reaction
50 mM Tris (pH 9.4), 300 mM NaCl
Buffer D
100 mM KHPO, 4 mM tetrabutylammonium hydrogen sulfate, pH 5.9
Buffer E
75% (v/v) buffer D, 25% (v/v) methanol
Equipment
Micropipettes with varying capacity (T-2, T-10, T-20, T-100, T-200, T-1000) (Tarsons, catalog numbers: 030000, 030010, 030020, 030030, 030040, 03005)
Spectrophotometer (Eppendorf, model: BioSpectrometer®)
HPLC system: Agilent 1200 HPLC (Agilent Technologies, model: Agilent 1200 HPLC) equipped with quaternary pump, autosampler, thermostated column compartment, with a UV detector
Äkta (Cytiva, former GE Health care, model: 29707638)
C-18 column (4.6 × 150 mm) (Agilent, model: Eclipse XDB-C-18)
Superose 12 10/300 Column (Cytiva, catalog number: GE17-5173-01)
Dry bath (Bio-Rad, catalog number: 1660563)
Refrigerated centrifuge (Eppendorf, model: centrifuge 5430)
Vortex shaker (Tarsons, model: SPINIXTM)
Biological safety cabinet (Thermo Scientific, model: 1300 Series Class II, Type A2)
Vacuum pump and assembly (Tarsons, model: ROCKYVACTM Vacuum Pump with assembly)
Water purification system (Millipore, model: Ultra-Pure Water Purification System)
Software and datasets
HPLC system software LC/CE Agilent ChemStation (B.04.02 SP1, 4/2010)
Graph Pad Prism 5.01 (5.01, 9/2007)
Procedure
DisA protein purification and C-di-AMP synthesis assay with DisA protein from Mycobacterium smegmatis
Purify the DisA protein as described in Gautam et al. [12]. The protein purification procedure is briefly described below.
Inoculate E. coli BL21 (DE3) carrying the pET28a plasmid with the disA gene in LB medium and grow overnight at 37 °C. Prepare secondary cultures by inoculating 1% of the primary culture and grow at 37 °C with shaking until the OD reaches 0.6.
Induce the secondary cultures by adding 1 mM isopropyl ß-D-thiogalactopyranoside (IPTG) and incubate for 3 h at 37 °C.
Harvest the cultures by centrifugation at 495× g; then, resuspend the pellet in lysis buffer [50 mM Tris-Cl (pH 7.9), 300 mM NaCl, and 1 mM phenylmethylsulfonyl fluoride (PMSF)].
Sonicate the cells and centrifuge at 1980× g to remove the cell debris.
Load the supernatant onto a Ni-NTA column to allow the protein to bind to the Ni-NTA beads.
Wash the column with 100 column volumes of wash buffer (50 mM Tris-Cl (pH 7.9), 300 mM NaCl, and 40 mM imidazole).
Elute the DisA protein using elution buffer containing 50 mM Tris-Cl (pH 7.9), 300 mM NaCl, and 300 mM imidazole. Analyze the protein by 10% SDS-PAGE.
Dialyze the purified DisA protein in Buffer A (see Recipes) for 12–16 h at 4 °C) as per the mentioned protocol [12].
Inject the dialyzed protein into a size exclusion chromatography (SEC) column to further purify the protein against buffer B (see Recipes).
Store the purified protein at -20 °C or -80 °C for future use.
Determine the concentration of the DisA protein (photometric measurement at 280 nm/BCA method).
Calculate the amount of DisA protein required to achieve a concentration of 1 μM in a 50 μL reaction.
In a 1.5 mL Eppendorf tube, set up a 50 μL reaction containing DisA protein (1 μM), 0.5 mM ATP, 5 mM MgCl in Buffer C (see Recipes).
Incubate the reaction at 37 °C for 4 h.
Stop the reaction by adding 10 mM EDTA to the reaction tube.
Centrifuge the reaction sample at 1,530× g for 30 min at 4 °C.
Collect the supernatant for HPLC analysis or store it at -20 °C for future use (Figure 1).
Figure 1. HPLC profiles of nucleotides and standard curve of c-di-AMP. A. HPLC profile of standard ATP. B. HPLC profile of standard C-di-AMP. C. HPLC profile of C-di-AMP synthesis reaction. D. Standard curve for c-di-AMP.
Generation of c-di-AMP standard curve
Prepare different concentrations of c-di-AMP or ATP (ranging from 100 to 500 μM) stock solutions in buffer C (see Recipes).
Use a reverse phase C-18 column to separate the nucleotides by using Buffer D (see Recipes) and Buffer E (see Recipes).
Filter and degas the HPLC buffers D and E using 0.22 μm filter paper.
Apply the following buffer gradient to separate the nucleotides at a flow rate of 0.7 mL/min:
0.0 min: 0% buffer E
2.5 min: 0% buffer E
5.0 min: 30% buffer E
10.0 min: 60% buffer E
14.0 min: 100% buffer E
21.0 min: 100% buffer E
22.0 min: 50% buffer E
23.0 min: 0% buffer E
Inject 20 μL of 500 μM pure c-di-AMP and then ATP separately for HPLC analysis.
After each sample injection, wash the HPLC column with buffer D for 10 min at a flow rate of 0.7 mL/min.
Pure c-di-AMP should show a peak at ~23.7 min, and ATP should show a peak at ~17.4 min.
After this preliminary validation, inject 20 μL of different concentrations (100–500 μM) of c-di-AMP into the HPLC and analyze using the previously stated gradient and buffers.
Use the chromatography data system software to obtain the peak area.
Prepare the standard curve by plotting the peak area under the curve (AUC) vs. the concentration of c-di-AMP used.
Analysis of the C-di-AMP synthesis assay reaction by HPLC
Use 20 μL of reaction sample for HPLC analysis.
Inject the reaction sample into the HPLC system using the gradient described above.
Observe for a peak at approximately ~23.7 min.
If a peak is observed at ~23.7 min, determine the area under the peak using the chromatography data system software.
Use the peak area to apply the standard curve and linear equation to determine the concentration (μM) of c-di-AMP formed in the reaction.
Collect the HPLC elution fractions around 23.7 min to further confirm the mass of the reaction product (See General note 2).
General notes and troubleshooting
The synthesis of c-di-AMP by DisA (a DAC domain-containing protein) highly depends on assay conditions like buffer compositions (NaCl/KCl), pH (5.4–9.4), temperature (35–40 °C), and incubation time (0–6 h). For Mycobacterium smegmatis, the highest activity was observed in Buffer C (50 mM Tris, 300 mM NaCl, pH 9.4). After 4 h of incubation at 37 °C, complete utilization of ATP was noted. When using DisA from different sources, activity optimization might be necessary to avoid multiple peaks (e.g., ATP or intermediate products).
We employ an earlier reported LC-MS method to analyze the HPLC eluted fractions from the DisA reactions (Burker Daltonics, Germany) as described by [14]. MS–MS analysis (negative or positive ion mode) of the eluted molecular masses is performed to verify the presence of c-di-AMP. The standard protocol involves LC–MS and MS–MS analysis using the HPLC gradient composition, as previously reported [15].
Validation of protocol
This protocol or parts of it has been used and validated in the following research article:
Ryjenkov DA et al. [10]. Cyclic diguanylate is a ubiquitous signalling molecule in bacteria: insights into biochemistry of the GGDEF protein domain. J Bacteriol. [Fig 2B, Fig 3B,C, Used for c-di-GMP analysis].
Christen M et al. [11]. Identification and characterization of a cyclic di-GMP-specific phosphodiesterase and its allosteric control by GTP. J Biol Chem. [Fig 2A, Used for c-di-GMP analysis].
Bai Y et al. [13]. Mycobacterium tuberculosis Rv3586 (DacA) is a diadenylate cyclase that converts ATP or ADP into c-di-AMP. PLoS One. [Fig 2A, Used for C-di-AMP analysis]
Acknowledgments
D.C. gratefully acknowledges the initial financial help through a grant from the Dept. of Biotechnology, Govt. Of India, through a grant number DBT/PR33123/MED/29/1497/2020.
Competing interests
The authors declare no competing interests.
References
Stülke, J. and Krüger, L. (2020). Cyclic di-AMP Signaling in Bacteria. Annu Rev Microbiol. 74(1): 159–179. https://doi.org/10.1146/annurev-micro-020518-115943
Corrigan, R. M. and Gründling, A. (2013). Cyclic di-AMP: another second messenger enters the fray. Nat Rev Microbiol. 11(8): 513–524. https://doi.org/10.1038/nrmicro3069
Zhang, L. and He, Z. G. (2013). Radiation-sensitive Gene A (RadA) Targets DisA, DNA Integrity Scanning Protein A, to Negatively Affect Cyclic Di-AMP Synthesis Activity in Mycobacterium smegmatis. J Biol Chem. 288(31): 22426–22436. https://doi.org/10.1074/jbc.m113.464883
Petchiappan, A., Mahapa, A. and Chatterji, D. (2020). Cyclic Dinucleotide Signaling in Mycobacteria. In: Chou, SH., Guiliani, N., Lee, V., Römling, U. (eds) Microbial Cyclic Di-Nucleotide Signaling. Springer, Cham. 3–25. https://doi.org/10.1007/978-3-030-33308-9_1
Sparks, I. L., Derbyshire, K. M., Jacobs, W. R. and Morita, Y. S. (2023). Mycobacterium smegmatis: The Vanguard of Mycobacterial Research. J Bacteriol. 205(1): e00337–22. https://doi.org/10.1128/jb.00337-22
Witte, G., Hartung, S., Büttner, K. and Hopfner, K. P. (2008). Structural Biochemistry of a Bacterial Checkpoint Protein Reveals Diadenylate Cyclase Activity Regulated by DNA Recombination Intermediates. Mol Cell. 30(2): 167–178. https://doi.org/10.1016/j.molcel.2008.02.020
Tang, Q., Luo, Y., Zheng, C., Yin, K., Ali, M. K., Li, X. and He, J. (2015). Functional Analysis of a c-di-AMP-specific Phosphodiesterase MsPDE from Mycobacterium smegmatis. Int J Biol Sci. 11(7): 813–824. https://doi.org/10.7150/ijbs.11797
He, Q., Wang, F., Liu, S., Zhu, D., Cong, H., Gao, F., Li, B., Wang, H., Lin, Z., Liao, J., et al. (2016). Structural and Biochemical Insight into the Mechanism of Rv2837c from Mycobacterium tuberculosis as a c-di-NMP Phosphodiesterase. J Biol Chem. 291(7): 3668–3681. https://doi.org/10.1074/jbc.m115.699801
Commichau, F. M., Heidemann, J. L., Ficner, R. and Stülke, J. (2019). Making and Breaking of an Essential Poison: the Cyclases and Phosphodiesterases That Produce and Degrade the Essential Second Messenger Cyclic di-AMP in Bacteria. J Bacteriol. 201(1): e00462–18. https://doi.org/10.1128/jb.00462-18
Ryjenkov, D. A., Tarutina, M., Moskvin, O. V. and Gomelsky, M. (2005). Cyclic Diguanylate Is a Ubiquitous Signaling Molecule in Bacteria: Insights into Biochemistry of the GGDEF Protein Domain. J Bacteriol. 187(5): 1792–1798. https://doi.org/10.1128/jb.187.5.1792-1798.2005
Christen, M., Christen, B., Folcher, M., Schauerte, A. and Jenal, U. (2005). Identification and Characterization of a Cyclic di-GMP-specific Phosphodiesterase and Its Allosteric Control by GTP. J Biol Chem. 280(35): 30829–30837. https://doi.org/10.1074/jbc.m504429200
Gautam, S., Mahapa, A., Yeramala, L., Gandhi, A., Krishnan, S., Kutti R., V. and Chatterji, D. (2023). Regulatory mechanisms of c‐di‐AMP synthase from Mycobacterium smegmatis revealed by a structure: Function analysis. Protein Sci. 32(3): e4568. https://doi.org/10.1002/pro.4568
Bai, Y., Yang, J., Zhou, X., Ding, X., Eisele, L. E. and Bai, G. (2012). Mycobacterium tuberculosis Rv3586 (DacA) Is a Diadenylate Cyclase That Converts ATP or ADP into c-di-AMP. PLoS One. 7(4): e35206. https://doi.org/10.1371/journal.pone.0035206
Bharati, B. K., Mukherjee, R. and Chatterji, D. (2018). Substrate-induced domain movement in a bifunctional protein, DcpA, regulates cyclic di-GMP turnover: Functional implications of a highly conserved motif. J Biol Chem. 293(36): 14065–14079. https://doi.org/10.1074/jbc.ra118.003917
Bharati, B. K., Sharma, I. M., Kasetty, S., Kumar, M., Mukherjee, R. and Chatterji, D. (2012). A full-length bifunctional protein involved in c-di-GMP turnover is required for long-term survival under nutrient starvation in Mycobacterium smegmatis. Microbiology (N Y). 158(6): 1415–1427. https://doi.org/10.1099/mic.0.053892-0
Article Information
Publication history
Received: Jul 30, 2024
Accepted: Oct 4, 2024
Available online: Nov 4, 2024
Published: Dec 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
How to cite
Category
Microbiology > Microbial biochemistry > Other compound
Biochemistry > Other compound
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Original Research Article:
The authors used this protocol in Cell Metabolism Feb 2024
Abstract
Sterol regulatory element binding proteins (SREBPs) are transcription factors that reside in the endoplasmic reticulum (ER) membrane as inactive precursors. To be active, SREBPs are translocated to the Golgi where the transcriptionally active N-terminus is cleaved and released to the nucleus to regulate gene expression. Nuclear SREBP levels can be determined by immunoblot analysis; however, this method can only determine the steady-state levels of nuclear SREBPs and does not capture the actual status of activation. The vesicle budding assay provides an alternative way to quantify the activation of SREBPs by monitoring the initiation of SREBP translocation from the ER to the Golgi through vesicles. Microsomal membranes isolated from the liver are incubated in a reaction buffer containing the necessary components to facilitate vesicle formation. Microsomal membranes and vesicles are isolated and SREBPs are quantified in each by immunoblot analysis. The amount of SREBPs found in the budded vesicles provides an assessment of the SREBP activation in the liver.
Key features
• This protocol describes a method to isolate budding vesicles from liver ER membranes
• The in vitro budding assay can be applied to investigate the movement of proteins from the ER to the Golgi
• This protocol was developed based on the procedures described previously with cultured cells [1–3]
Keywords: Endoplasmic reticulum (ER) Golgi complex (Golgi) Microsomal isolation Cytosol preparation Budding assay Sterol regulatory element binding protein (SREBP)
Graphical overview
Background
The transport of proteins from the endoplasmic reticulum (ER) to the Golgi complex is required for additional protein modifications and/or protein secretion. Small transport vesicles that bud from mammalian ER and fuse to Golgi were reported in the late 1980s [4] and further characterized thereafter [2,3,5]. Sterol regulatory element binding proteins (SREBPs) are a family of membrane-bound transcription factors that regulate the expression of genes involved in cholesterol and fatty acid homeostasis [6–8], which utilize small vesicles for transport from the ER to the Golgi for activation [1]. Newly synthesized SREBP proteins are embedded in the ER membrane, forming a complex with SREBP cleavage activating protein (SCAP), a protein that serves as a sterol sensor and escort protein. The process of SREBP activation requires the membrane-bound SREBP/SCAP complex to move from the ER to Golgi, where two proteases reside, which cleave and free the transcriptionally active N-terminal portion of SREBPs from the membrane. The N-terminal SREBPs then enter the nucleus and regulate the transcription of downstream genes [8].
Immunoblot analysis is routinely used to evaluate the degree of SREBP activation. The molecular weight of the cleaved mature/nuclear form of SREBPs is ~68 kD, while the molecular weight of the membrane-bound precursor SREBP is ~125 kD. By quantifying the amount of cleaved nuclear SREBPs, an estimation of SREBP activation can be achieved. However, nuclear SREBP levels represent a steady-state measurement of the combined results of both protein activation and degradation. In a recent study, we developed a new method to determine SREBP activation by measuring the full-length SREBPs in budding vesicles of mouse liver. Although the in vitro budding reaction might have the limitation of not reflecting the in vivo status, this method measures the initiation of SREBP transportation from the ER to Golgi for cleavage activation. Compared to measuring the cleaved nuclear form SREBPs, evaluation of SREBP activation by measuring the SREBPs in the budding vesicles eliminates the potential effects of degradation of nuclear SREBPs.
Materials and reagents
Biological materials
Note: The general microsomal budding assay is not dependent on the animal or diet used.
Human SREBP-1c transgenic rat
High carbohydrate diet (MP Biomedicals, catalog number: 960238)
Reagents
HEPES (Sigma-Aldrich, catalog number: H3375)
D-Sorbitol (Sigma-Aldrich, catalog number: S6021)
Potassium acetate (Sigma-Aldrich, catalog number: P1190)
Magnesium acetate tetrahydrate (Sigma-Aldrich, catalog number: M5661)
0.5 M EGTA-KOH, pH 8.0 (Fisher Scientific, catalog number: 50-997-744)
1 M Tris-Cl, pH 6.8 (Fisher Scientific, catalog number: 50-843-263)
Dithiothreitol (DTT) (Sigma-Aldrich, catalog number: D9779)
Sodium dodecyl sulfate (SDS) (Sigma-Aldrich, catalog number: 75746)
Bromophenol blue (Sigma-Aldrich, catalog number: B0126)
Glycerol (Sigma-Aldrich, catalog number: G5516)
Trizma base (Sigma-Aldrich, catalog number: T1503)
Protease inhibitor cocktail (Sigma-Aldrich, catalog number: P8340)
0.9% Sodium chloride irrigation (Saline) (Baxter, catalog number: BX-2F7124)
Isoflurane (Piramal Critical Care, catalog number: 33794-013-25)
Creatine phosphate (Millipore Sigma, catalog number: 10621714001)
ATP (Sigma-Aldrich, catalog number: G6419)
GTP (Sigma-Aldrich, catalog number: G8877)
Creatine kinase (Millipore Sigma, catalog number: 10127566001)
Anti-HA antibody (Cell Signaling, catalog number: 3724)
Anti-Calnexin antibody (Enzo, catalog number: ADI-SPA-860-F)
Anti-ERGIC-53 antibody (Abcam, catalog number: ab125006)
Goat anti-rabbit IgG, F(ab’)2 (Jackson Immuno, catalog number: 115-035-072)
PierceTM BCA Protein Assay kit (Thermo Fisher, catalog number: A65453)
SuperSignal West Pico chemiluminescent substrate (Thermo Fisher, catalog number: 23225)
Solutions
Buffer 1 (see Recipes)
Buffer 2 (see Recipes)
5× SDS-loading buffer (see Recipes)
20× budding reaction supplement (see Recipes)
Recipes
pH values of buffers are all adjusted at room temperature.
Buffer 1
Reagent Final concentration Quantity or Volume
1 M HEPES-KOH, pH 7.2 10 mM 500 μL
2 M Sorbitol 250 mM 6.25 mL
1 M KOAc 10 mM 500 µL
1 M Mg(OAc)2 1.5 mM 75 µL
Total n/a Adjust pH to 7.2 with KOH, then add H2O to 50 mL
Can be stored at 4 °C for up to two months.
Buffer 2
Reagent Final concentration Volume
1 M HEPES-KOH, pH 7.2 50 mM 2.5 mL
2 M Sorbitol 250 mM 6.25 mL
1 M KOAc 70 mM 3.5 mL
1 M Mg(OAc)2 2.5 mM 125 µL
0.5 M EGTA-KOH, pH 8.0 5 mM 500 µL
Total n/a Adjust pH to 7.2 with KOH, then add H2O to 50 mL
Can be stored at 4 °C for up to two months.
5× SDS-loading buffer
Reagent Final concentration Volume
SDS 15% 15 g
Bromophenol blue 0.02% 20 mg
Glycerol 25% 25 mL
1 M Tris-Cl, pH 6.8 0.15 M 15 mL
Total n/a Add H2O to 100 mL
Add 62.5 μL of β-mercaptoethanol for each 500 μL of loading buffer freshly before use.
20× budding reaction supplement
The following reagents should be freshly prepared individually. Prepare the 20× budding reaction supplement by mixing equal volumes of each reagent immediately before carrying out the budding reaction.
Note: The molecular weight or specific activity of each reagent will be different depending on the source and lot number of the reagents. The quantity needs to be calculated based on the information from the vendor.
Reagent 20× concentration Quantity Final volume in H2O
Creatine phosphate 200 mM 26.2 mg 100 µL
ATP 30 mM 6.61 mg 100 µL
GTP 10 mM 2.09 mg 100 µL
Creatine kinase 80 U/mL 1 mg 10 mL
Laboratory supplies
18 G catheter (BD, catalog number: 382644)
Protein LoBind tubes (Eppendorf, catalog number: 022431081)
Polycarbonate centrifuge tubes (for AT3 rotor) (Beckman Coulter, catalog number: 343776)
Polycarbonate centrifuge tubes (for AT6 rotor) (Beckman Coulter, catalog number: 362305)
Equipment
Tabletop centrifuge (Eppendorf, model: Centrifuge 5424)
37 °C incubator (ThermoMixer, Eppendorf, model: F1.5 was used in this experiment)
Ultracentrifuge (Thermo Scientific, model: Sorvall MX120)
S100-AT6 fixed angle rotor (Thermo Scientific, catalog number: 45588)
S120-AT3 fixed angle rotor (Thermo Scientific, catalog number: 45584)
Peristaltic pump (Amersham Biosciences, model: Pump P-1)
Dounce tissue homogenizer-2 mL (DWK life sciences, catalog number: 885300-002)
Dounce tissue homogenizer-7 mL (DWK life sciences, catalog number: 885300-007)
Procedure
Preparation of cytosol from rat liver
Note: The rat used for cytosol preparation can be different from the animal used for microsomal preparation in section B. Cytosol can be prepared and aliquoted at -80 °C for long-term storage.
Fill the liver perfusion tube with ~100 mL of 0.9% saline and connect one end of the bubble trap tubular to the peristaltic pump and the other end to an 18 G catheter.
Euthanize the rat in an anesthetization jar filled with isoflurane.
Place the rat on a procedure pad, expose the abdominal cavity, and move the liver to expose the portal vein and the inferior vena cava (IVC).
Use an 18 G catheter to cannulate the portal vein; then, cut the IVC using scissors, start the perfusion pump at a rate of ~5 mL/min, and perfuse for 10 min (Figure 1A).
Figure 1. Preparation of cytosol from rat liver. A. Liver perfusion with 0.9% saline. The blue arrow indicates the portal vein location where the catheter was inserted. The yellow arrow shows the location where the IVC was cut. B–E. Samples after each centrifugation in the cytosol preparation steps. Lipid fractions (top) and pellets were discarded after each centrifugation. The animal experiments were performed with the approval of the Institutional Animal Care and Use Committee at the University of Texas Southwestern Medical Center.
Remove the perfused liver and homogenize 2 g of liver in 5 mL of ice-cold buffer 2 supplemented with 1mM DTT using a 7 mL glass Dounce tissue homogenizer.
Centrifuge the homogenate at 4 °C as detailed below.
1,000 RCF for 10 min with a tabletop centrifuge. Remove the lipid fraction that floats on the supernatant with a pipette and only collect the clear fraction between the lipid layer and the pellet (Figure 1B). Transfer the clear supernatant to a polycarbonate centrifuge tube.
20,000 RCF for 20 min in an AT6 rotor using an ultracentrifuge machine; transfer the clear supernatant to a new polycarbonate centrifuge tube (Figure 1C).
186,000 RCF for 1 h in an AT6 rotor using an ultracentrifuge machine; transfer the clear supernatant to a new polycarbonate centrifuge tube (Figure 1D).
186,000 RCF for 45 min in an AT6 rotor using an ultracentrifuge machine (Figure 1E).
Collect the supernatant from the last spin (cytosol) and determine the protein concentration using a BCA Protein Assay kit. Aliquot the cytosol to individual tubes and store at -80 °C for future use.
Microsomal membrane isolation from liver
This protocol has been validated with mouse and rat livers.
Homogenize 200 mg of fresh liver in 1 mL of ice-cold buffer 1 supplemented with a protease inhibitor cocktail (100×) using a 2 mL Dounce homogenizer.
Centrifuge the liver homogenate at 1,500 RCF for 5 min in a tabletop centrifuge at 4 °C. Transfer the supernatant to a new tube and discard the pellet (Figure 2A).
Figure 2. Samples after centrifugation during microsomal membrane preparations. Panels A–E show the outcomes after centrifugation in steps B2, B4, B5, B7, and C3 respectively.
Centrifuge the supernatant from step B2 at 1,500 RCF for another 5 min at 4 °C.
Transfer the supernatant to a 1.5 mL protein low-bind tube. Avoid disturbing the remaining small pellet at the bottom (Figure 2B).
Centrifuge the supernatant at 16,000 RCF for 10 min in a tabletop centrifuge at 4 °C and remove and discard the supernatant (Figure 2C).
Resuspend the pellet in 400 µL of ice-cold buffer 2 supplemented with a protease inhibitor cocktail (100×) and transfer to a new protein low-bind tube.
Centrifuge the resuspended pellet at 16,000 RCF for 3 min in a tabletop centrifuge at 4 °C and remove and discard the supernatant (Figure 2D).
Resuspend the pellet (microsomal membranes) in 70 μL of ice-cold buffer 2 supplemented with the protease inhibitor cocktail (100×).
Determine the microsomal membrane protein concentration using the BCA Protein Assay kit.
In vitro budding reaction
For each reaction, add 80 μg of microsomal protein, 4 μL of the 20× budding reaction supplement, 600 μg of liver cytosol, and adjust the final reaction volume to 80 μL with buffer 2. Mix the reaction by pipetting up and down 10 times.
Incubate the reaction at 37 °C for 20 min, then transfer to ice for 5 min.
Centrifuge the reaction from step C2 at 16,000 RCF in a tabletop centrifuge for 3 min at 4 °C (Figure 2E).
Budding vesicle collection:
Transfer 65 μL of the supernatant from step C3 to a polycarbonate centrifuge tube and centrifuge at 137,000 RCF for 30 min at 4 °C in an AT3 rotor using an ultracentrifuge machine to isolate the formed vesicles.
Remove the supernatant, resuspend the pellet in 30 μL of buffer 2 supplemented with 5× SDS-loading buffer, and incubate at 55 °C for 20 min. Following the incubation, load the entire sample to an 8% SDS-PAGE for immunoblot analysis of proteins of interest that reside in the isolated budding vesicles.
Microsomal membrane collection: Resuspend the pellet (microsomal membrane) from step C3 in 60 μL of buffer 2 supplemented with 5× SDS-loading buffer and incubate at 55 °C for 20 min. Following the incubation, load 15 μL of the sample onto an 8% SDS-PAGE for immunoblot analysis.
Note: You might only see a very tiny pellet following the centrifugation in step C5. Mark the tube position in the rotor to help locate the pellet after centrifugation.
Validation of protocol
This protocol has been used and validated in the following research article:
Rong et al. [8]. DGAT2 inhibition blocks SREBP-1 cleavage and improves hepatic steatosis by increasing phosphatidylethanolamine in the ER. Cell Metabolism (Figure S7, panel D).
This protocol was developed based on previous research articles that used the budding assay in cultured cells:
Rexach et al. [2]. Distinct biochemical requirements for the budding, targeting, and fusion of ER-derived transport vesicles. The Journal of Cell Biology.
Rowe et al. [3]. COPII vesicles derived from mammalian endoplasmic reticulum microsomes recruit COPI. The Journal of Cell Biology.
Nohturfft et al. [1]. Regulated step in cholesterol feedback localized to budding of SCAP from ER membranes. Cell.
We also validated this protocol using transgenic rats that constitutively express HA-tagged human full-length (precursor) SREBP-1c under the control of the apoE promoter (TghSREBP-1c) [10] (Figure 3). The rats were either fasted for 24 h (fast) (low levels of SREBP-1 activation) or for 18 h and then re-fed a high carbohydrate diet for 6 h (re-fed) (high levels of SREBP-1 activation) before microsome/vesicle isolation to achieve the maximal differences in SREBP-1 activation. The isolated budding vesicles were validated by immunoblot with anti-calnexin and anti-ERGIC-53 antibodies. Calnexin, a resident ER protein, was not detectable in the budding vesicles (Figure 3, lanes 3 and 4), while ERGIC-53, a protein that migrates together with budding vesicles [11,12], was detected in both microsomal membranes and budding vesicles, indicating the successful isolation of ER membrane–derived budding vesicles without microsomal contamination. The experiment also showed that SREBP-1 is enriched in the vesicles budded from microsomal membranes of re-fed rats (Figure 3, lane 4 vs. lane 3), while ERGIC-53, serving as a loading control for budding vesicles, is similar between fast and re-fed groups, indicating that re-feeding the high carbohydrate diet increased SREBP-1 activation.
Figure 3. Immunoblot analysis of sterol regulatory element binding protein 1 (SREBP-1) in microsomal membranes and budding vesicles. TghSREBP-1c rats were fasted for 24 h (fast) or for 18 h and then re-fed a high carbohydrate diet for 6 h (re-fed). Liver microsomal membranes and budding vesicles were prepared for immunoblot analysis. Lane 1 represents microsomal membranes from fasted rats; lane 2 is microsomal membranes from re-fed rats; lane 3 is budding vesicles from fasted rat microsomes; lane 4 is budding vesicles from re-fed rat microsomes. The full length of transgenic human SREBP-1c protein (molecular weight ~ 125 kD) was detected using an anti-HA antibody. Calnexin (molecular weight ~ 95 kD) is a loading marker for microsome membranes. ERGIC-53 (molecular weight ~ 53 kD) is a protein associated with the vesicles during the budding process.
Acknowledgments
We thank Dr. Jay D. Horton for critical reading of the manuscript. This work was supported by NIH grants 5P01HL160487 and 5P30DK127984. This protocol was developed based on previous research articles that used cultured cells for budding assay [1–3] and has been used and validated in Cell Metabolism (2024), DOI: 10.1016/j.cmet.2024.01.011 [8].
Competing interests
The authors have no competing interests to claim.
Ethical considerations
The animal experiments were performed with the approval of the Institutional Animal Care and Use Committee at the University of Texas Southwestern Medical Center.
References
Nohturfft, A., Yabe, D., Goldstein, J. L., Brown, M. S. and Espenshade, P. J. (2000). Regulated step in cholesterol feedback localized to budding of SCAP from ER membranes. Cell. 102(3): 315–323. https://doi.org/10.1016/s0092-8674(00)00037-4.
Rexach, M. F. and Schekman, R. W. (1991). Distinct biochemical requirements for the budding, targeting, and fusion of ER-derived transport vesicles. J Cell Biol. 114(2): 219–229. https://doi.org/10.1083/jcb.114.2.219.
Rowe, T., Aridor, M., McCaffery, J. M., Plutner, H., Nuoffer, C. and Balch, W. E. (1996). COPII vesicles derived from mammalian endoplasmic reticulum microsomes recruit COPI. J Cell Biol. 135(4): 895–911. https://doi.org/10.1083/jcb.135.4.895.
Paulik, M., Nowack, D. D. and Morre, D. J. (1988). Isolation of a vesicular intermediate in the cell-free transfer of membrane from transitional elements of the endoplasmic reticulum to Golgi apparatus cisternae of rat liver. J Biol Chem. 263(33): 17738–17748.
Schekman, R. and Orci, L. (1996). Coat proteins and vesicle budding. Science. 271(5255): 1526–1533. https://doi.org/10.1126/science.271.5255.1526.
Shimomura, I., Shimano, H., Korn, B. S., Bashmakov, Y. and Horton, J. D. (1998). Nuclear sterol regulatory element-binding proteins activate genes responsible for the entire program of unsaturated fatty acid biosynthesis in transgenic mouse liver. J Biol Chem. 273(52): 35299–35306. https://doi.org/10.1074/jbc.273.52.35299.
Horton, J. D., Goldstein, J. L. and Brown, M. S. (2002). SREBPs: activators of the complete program of cholesterol and fatty acid synthesis in the liver. J Clin Invest. 109(9): 1125–1131. https://doi.org/10.1172/JCI15593.
Rong, S., Xia, M., Vale, G., Wang, S., Kim, C. W., Li, S., McDonald, J. G., Radhakrishnan, A. and Horton, J. D. (2024). DGAT2 inhibition blocks SREBP-1 cleavage and improves hepatic steatosis by increasing phosphatidylethanolamine in the ER. Cell Metab. 36(3): 617–629 e617. https://doi.org/10.1016/j.cmet.2024.01.011.
Brown, M. S. and Goldstein, J. L. (2009). Cholesterol feedback: from Schoenheimer's bottle to Scap's MELADL. J Lipid Res. 50 Suppl: S15–27. https://doi.org/10.1194/jlr.R800054-JLR200.
Owen, J. L., Zhang, Y., Bae, S. H., Farooqi, M. S., Liang, G., Hammer, R. E., Goldstein, J. L. and Brown, M. S. (2012). Insulin stimulation of SREBP-1c processing in transgenic rat hepatocytes requires p70 S6-kinase. Proc Natl Acad Sci USA. 109(40): 16184–16189. https://doi.org/10.1073/pnas.1213343109.
Klumperman, J., Schweizer, A., Clausen, H., Tang, B. L., Hong, W., Oorschot, V. and Hauri, H. P. (1998). The recycling pathway of protein ERGIC-53 and dynamics of the ER-Golgi intermediate compartment. J Cell Sci. 111 (Pt 22): 3411–3425. https://doi.org/10.1242/jcs.111.22.3411.
Appenzeller, C., Andersson, H., Kappeler, F. and Hauri, H. P. (1999). The lectin ERGIC-53 is a cargo transport receptor for glycoproteins. Nat Cell Biol. 1(6): 330–334. https://doi.org/10.1038/14020.
Article Information
Publication history
Received: Sep 18, 2024
Accepted: Oct 21, 2024
Available online: Nov 4, 2024
Published: Dec 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Category
Molecular Biology > Protein > Activity
Cell Biology > Cell-based analysis > Protein maturation
Cell and Molecular Biology
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Immunofluorescence for Detection of TOR Kinase Activity In Situ in Photosynthetic Organisms
AL Ana P. Lando §
MM María A. De Marco
AC Andrea C. Cumino
GM Giselle M. A. Martínez-Noël §
(§Technical contact: [email protected]; [email protected])
Published: Vol 14, Iss 24, Dec 20, 2024
DOI: 10.21769/BioProtoc.5140 Views: 316
Reviewed by: Shuhei OtaMalgorzata Lichocka Anonymous reviewer(s)
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Cited by
Original Research Article:
The authors used this protocol in Planta Oct 2024
Abstract
The target of rapamycin (TOR) is a central hub kinase that promotes growth and development in all eukaryote cells. TOR induces protein synthesis through the phosphorylation of the S6 kinase (S6K), which, in turn, phosphorylates ribosomal S6 protein (RPS6) increasing this anabolic process. Therefore, S6K and RPS6 phosphorylation are generally used as readouts of TOR activity. Protein phosphorylation levels are measured by a western blot (WB) technique using an antibody against one specific phosphosite in cell extracts. However, at the tissue/cell-specific level, there is a huge gap in plants due to the lack of alternative techniques for the evaluation of TOR activity as there are for other organisms such as mammals. Here, we describe an in vivo protocol to detect S6K phosphorylation in tissues/cells of model photosynthetic organisms such as Arabidopsis thaliana and Chlamydomonas reinhardtii. Our proposed method consists of the immunolocalization of a phosphorylated target of TOR kinase using a fluorescent secondary antibody by confocal microscopy. The protocol involves four main steps: tissue/cell fixation, permeabilization, and incubation with primary and secondary antibodies. It is an easy technique that allows handling different samples at the same time. In addition, different ultrastructural cell markers can also be used, such as for nucleus and cell wall detection, allowing a detailed analysis of cell morphology. To our knowledge, this is the first protocol to detect TOR activity in situ in photosynthetic organisms; we consider that it will pave the research on the TOR kinase, opening new possibilities to better understand its complex signaling.
Key features
• The protocol is an easy and non-destructive method to detect S6K phosphorylation at the cellular level for plants and algae.
• First method for in situ immunolocalization of target proteins of TOR kinase in photosynthetic organisms.
Keywords: Arabidopsis Chlamydomonas Confocal microscopy Immunofluorescence in situ S6K TOR activity
Graphical overview
Background
The target of rapamycin (TOR) kinase pathway is an ancestral signaling pathway that integrates nutrient information with translational control and growth regulation [1,2]. This conserved signaling pathway includes the S6K and its direct target, the 40S ribosomal protein S6 (RPS6) [3]. Therefore, the measurement of S6K and RPS6 phosphorylation levels through the WB technique is used as a reliable readout of TOR activity in eukaryotes including photosynthetic organisms [4,5]. The method is generally accurate in estimating TOR activity but requires cell disruption and does not permit subcellular localization or monitoring tissue specificities. Other techniques, such as immunofluorescence, have been described for mammals; however, in plants and algae, there is still a need to develop protocols for detecting TOR activity at the tissue/cell-specific level. Immunolocalization approaches can provide a great deal of information about the subcellular localization and dynamics of many plant proteins, even in vivo [6], which is particularly important when we have multi-level regulation, as is the case with the TOR kinase. We have developed, analyzed, and validated an immunofluorescence protocol for detecting TOR activity through S6K phosphorylation in situ in model photosynthetic organisms (Arabidopsis and Chlamydomonas). The main advantages of this method include its applicability to both plants and algae, its simplicity as it does not require tissue sectioning, its reproducibility due to Alexa's fluorophore stability, and the visualization in vivo due to the non-destruction of tissues/cells. Some minor limitations of the protocol include the restriction of its application to young tissues of plants, some reagents being expensive such as fluorescent antibodies, and the need for specialized personnel for the management of the confocal microscope.
This protocol will allow progress in the studies on TOR signaling at the subcellular level and tissue specificities, which are known to be essential for this pathway and where information is lacking in photosynthetic organisms, probably due to technical limitations.
To our knowledge, this is the first protocol for the analysis of TOR activity at the cellular level in plants and algae.
Materials and reagents
Biological materials
Arabidopsis thaliana Columbia ecotype (Col-0) wild-type (WT), young seedlings (5 days old)
Chlamydomonas reinhardtii CC125 (137c, mt + nit1 nit2), 15 mL of OD750nm: 0.6–0.9
Reagents
Double-distilled water (ddH2O)
Sodium phosphate dibasic (Na2HPO4) (Sigma, catalog number: 71496)
Potassium dihydrogen phosphate (KH2PO4) (Sigma, catalog number: 60220-M)
Sodium chloride (NaCl) (Anedra, catalog number: AN00716909)
Orthophosphoric acid (H3PO4) (Sigma, catalog number: 695017)
Paraformaldehyde (Agar Scientific, catalog number: R1018)
Sodium hydroxide (NaOH) (Fluka, catalog number: 71690)
Bovine serum albumin (BSA) 10 mg/mL (Promega, catalog number: R396D)
Triton X-100 (Sigma, catalog number: 9036-19-5)
Primary polyclonal antibody p-p70 S6K1 α Thr 389 (Santa Cruz, catalog number: sc-11759)
Secondary antibody anti-IgG-Alexa 488 (Invitrogen, catalog number: A11008)
Propidium iodide (PI) (Invitrogen, catalog number: P1304MP)
4',6-Diamidino-2-phenylindole dihydrochloride (DAPI) (Sigma, catalog number: D9564)
Solutions
Paraformaldehyde (PFA) 4% (see Recipes)
Phosphate buffered saline (PBS) 5× (see Recipes)
Permeabilization and blocking buffer (PB) (see Recipes)
Recipes
PFA 4%
Weigh 4 g of PFA and place in 35 mL of double-distilled water at 60 °C, add 30–50 μL of 10 N NaOH, and leave to stir on a hot plate (until the PFA dissolves well).
Add 20 mL of 5× PBS.
Check pH; it should be between 7.2 and 7.8.
Bring up to 100 mL with ddH2O.
Split into Falcon tubes and store at -20 °C for up to 3 months and protected from light.
Caution: Highly toxic. Wear protective gloves and work in the fume hood when handling the powder.
PBS (5×) (for 1 L)
Dissolve 3.62 g of Na2HPO4 (FW 141.96), 1.05 g of KH2PO4 (FW 136.1), and 38.25 g of NaCl (FW 58.44) in 800 mL of ddH2O. Adjust the pH to 7.4 with H3PO4 and then add ddH2O to the final volume. Dispense the solution into aliquots and sterilize them by autoclaving. The work solution used is 1× (final concentration 8 mM Na2HPO4, 2 mM KH2PO4, and 140 mM NaCl).
Permeabilization and blocking buffer (PB) (for 1.5 mL of PB)
Add 150 μL of 1% BSA, 45 μL of 10% Triton X-100, and 1,305 μL of 1× PBS (final concentration 1× PBS, 0.1% BSA, and 0.3% Triton X-100).
Equipment
Confocal microscope (e.g., Nikon, model: Eclipse C1 Plus)
Vacuum pump with a desiccator
Software and datasets
Confocal microscope software (e.g., EZ-C1 software Nikon)
NIH ImageJ software 1.47 for Windows (https://imagej.net/ij/)
Procedure
Tissue fixation (duration 4–6 h)
Remove Arabidopsis or Chlamydomonas samples from the growth medium and wash with 1× PBS.
Add 200 μL of 4% PFA to each tube (must cover the biological material).
Sample incubation
For algae: Incubate for 6 h at 4 °C.
For plant: Cut the root, place it in a microcentrifuge tube, and incubate the tissue in a vacuum desiccator for 6 h at room temperature (approximately 20 °C) to ensure proper tissue penetration.
Note: Leaves could also be used for the analysis (after removal of chlorophyll and other pigments).
Permeabilization and blocking (duration 24 h)
Remove the PFA solution and wash the material with 1× PBS at RT.
Add ~70 μL of PB to each tube (must cover the biological material) and incubate for 24 h at 4 °C.
Primary antibody incubation (duration 4 d)
Remove PB from all tubes (except the negative control, in which you should change the PB solution).
Add the antibody at a dilution of 1/50 in PB to a final volume of 50 μL per tube (except the negative control, in which you should change the PB solution).
Incubate for 4 days at 4 °C (move the solution every 1 or 2 days).
Wash the material with 1× PBS (wash, remove, and add back).
Incubate for 24 h at 4 °C.
Note: In the case of plants, primary antibody can be recovered for another use.
Secondary antibody incubation (duration 2 d)
Remove the PBS solution.
Add the anti-IgG-Alexa 488 antibody at a dilution of 1/250 in PB to a final volume of 50 μL per tube.
Incubate for 24 h at 4 °C.
Wash the material with 1× PBS (wash, remove, and add back).
Incubate for 24 h at 4 °C.
Note: Always protect the samples from light.
Nucleus staining (optional)
Nucleus labeling can be useful to localize the protein of interest at subcellular, cellular, and tissue level.
For propidium iodide (PI) staining:
Remove the PBS solution.
Add PI fluorophore to the samples at a dilution of 1/1,000 in 1× PBS to a final volume of 50 μL per tube (protect from light).
Incubate for 5 min at room temperature.
Wash the material four times with 1× PBS (wash, remove, and add back).
Analyze under a confocal microscope.
For DAPI staining:
Remove the PBS solution.
Add DAPI fluorophore to the samples at a dilution of 1/1,000 in 1× PBS to a final volume of 50 μL per tube (protect from light).
Incubate for 30 min for Arabidopsis or 5 min for Chlamydomonas at room temperature.
Wash the material three times with 1× PBS (wash, remove, and add back).
Analyze under a confocal microscope.
Data analysis
Sample images were obtained with C1 confocal laser using the following settings: green fluorescence intensity; excitation/emission wavelength = 488/561 nm. Visualization was done with the Super Fluor 40.0×/1.30/0.22 oil spring–loaded objective, and images were processed with the NIH ImageJ software. The positive signal appeared as bright spots in immunolocalization using a P-S6K antibody as was previously reported in rat and mouse tissues and human cell lines [7]. For Arabidopsis, at least eight roots were visualized (Figures 1A and 2A). For Chlamydomonas, eight coverslips were taken from each biological replicate (Figure 3A).
Figure 1. Immunolocalization of P-S6K in the root tip of five-day-old Arabidopsis seedlings. Confocal images of in situ immunofluorescence assays performed with the primary antibody p-p70 S6 kinase α Thr 389 and the secondary antibody anti-IgG-Alexa 488 (green fluorescence). P-S6K detection in the root tip zone of seedlings grown in control Murashige and Skoog (MS) medium (A) and treated with 2 μM AZD-8055 (TOR inhibitor) (B) for 48 h in darkness. Negative control consisted of the omission of primary antibody (C). Scale bars: 20 μm. Insets: Brightfield photos.
Figure 2. Immunolocalization of P-S6K in the elongation and differentiation zone of the root of five-day-old Arabidopsis seedlings. Confocal images of in situ immunofluorescence assays performed with the antibody p-p70 S6 kinase α Thr 389 and the secondary antibody anti-IgG-Alexa 488 (green fluorescence). P-S6K detection in the elongation and differentiation zone of the root of seedlings grown in control Murashige and Skoog (MS) medium (A) and treated with 2 μM AZD-8055 inhibitor (B) for 48 h in darkness. Negative control consisted of the omission of primary antibody (C). Scale bars: 50 μm. Insets: Brightfield photos.
Figure 3. Immunolocalization of P-S6K from Chlamydomonas. Confocal images of in situ immunofluorescence assays performed with the antibody p-p70 S6 kinase α Thr 389 and the secondary antibody anti-IgG-Alexa 488 (green fluorescence). P-S6K detection in Chlamydomonas grown in control medium TAP (A) and treated with 700 nM AZD-8055 inhibitor (B) for 24 h. Negative control consisted of the omission of primary antibody (C). Scale bars: 10 μm. Insets: Brightfield photos.
Validation of protocol
To validate the observed fluorescence pattern of the protein under study, each immunolocalization experiment must include the correct controls. In our case, we used an ATP-competitive TOR inhibitor (AZD-8055 Cayman, catalog number: 16978) to evaluate the specificity of the protocol. The AZD treatment considerably reduced the fluorescence intensity in both Arabidopsis and Chlamydomonas samples, corroborating that the signal detected is specifically due to the TOR activity (Figures 1B, 2B, and 3B). Besides, a sample without the addition of the primary antibody was used to determine the amount of background signal caused by the secondary antibody (Figures 1C, 2C, and 3C). This negative control did not show fluorescence, demonstrating that the technique has a low background. We also corroborated this methodology in other plant tissues such as lateral and hair roots (Figures S1 and S2).
General notes and troubleshooting
Troubleshooting
Regarding plant samples, the main limitation of this protocol is its restricted applicability to young and permeable tissues. To ensure the fixation of the Arabidopsis samples, it is mandatory to make a cut in the fresh tissue, and it is critical to perform the vacuum step, also for the root tip. While this step is not essential for other plant tissues and algae samples, it improves the methodology. In the case of cotyledons and young leaves, special care must be taken with interferences caused by chlorophyll, which must be removed using solvents. For microalgae, the material quantity is a crucial factor (~2–4 mg dry weight): it must be enough to be processed (taking into account material losses due to washing) and not be in excess to avoid interferences with cell fixation and permeabilization.
Acknowledgments
This work was supported by the National Agency for Promotion of Science and Technology (ANPCyT, PICT2019-2118), National Scientific and Technical Research Council (CONICET, PIP-11220200101701CO) and National University of Mar del Plata (UNMdP, EXA1142/23). This protocol was adapted and modified from Loos et al. [8] (doi: 10.1128/AAC.01808-19, PMID: 32540980).
Competing interests
The authors declare that they have no competing interests.
References
Pacheco, J. M., Canal, M. V., Pereyra, C. M., Welchen, E., Martínez-Noël, G. M. A. and Estevez, J. M. (2021). The tip of the iceberg: emerging roles of TORC1, and its regulatory functions in plant cells. J Exp Bot. 72(11): 4085–4101.
Artins, A., Martins, M. C. M., Meyer, C., Fernie, A. R. and Caldana, C. (2024). Sensing and regulation of C and N metabolism – novel features and mechanisms of the TOR and SnRK1 signaling pathways. Plant J. 118(5): 1268–1280.
Van Leene, J., Han, C., Gadeyne, A., Eeckhout, D., Matthijs, C., Cannoot, B., De Winne, N., Persiau, G., Van De Slijke, E., Van de Cotte, B., et al. (2019). Capturing the phosphorylation and protein interaction landscape of the plant TOR kinase. Nat Plants. 5(3): 316–327.
Dobrenel, T., Mancera-Martínez, E., Forzani, C., Azzopardi, M., Davanture, M., Moreau, M., Schepetilnikov, M., Chicher, J., Langella, O., Zivy, M., et al. (2016). The Arabidopsis TOR Kinase Specifically Regulates the Expression of Nuclear Genes Coding for Plastidic Ribosomal Proteins and the Phosphorylation of the Cytosolic Ribosomal Protein S6. Front Plant Sci. 7: e01611.
Upadhyaya, S., Agrawal, S., Gorakshakar, A. and Rao, B. J. (2020). TOR kinase activity in Chlamydomonas reinhardtii is modulated by cellular metabolic states. FEBS Lett. 594(19): 3122–3141.
Sauer, M., Paciorek, T., Benková, E. and Friml, J. (2006). Immunocytochemical techniques for whole-mount in situ protein localization in plants. Nat Protoc. 1(1): 98–103.
Schmidt, T., Wahl, P., Wüthrich, R. P., Vogetseder, A., Picard, N., Kaissling, B. and Le Hir, M. (2006). Immunolocalization of phospho-S6 kinases: a new way to detect mitosis in tissue sections and in cell culture. Histochem Cell Biol. 127(2): 123–129.
Loos, J. A., Dávila, V. A., Brehm, K. and Cumino, A. C. (2020). Metformin Suppresses Development of the Echinococcus multilocularis Larval Stage by Targeting the TOR Pathway. Antimicrob Agents Chemother. 64(9): e01808–19.
Supplementary information
The following supporting information can be downloaded here:
Figure S1. Immunolocalization of P-S6K in the root tip of five-day-old Arabidopsis seedlings
Figure S2. Immunolocalization of P-S6K in the root of five-day-old Arabidopsis seedlings
Article Information
Publication history
Received: Jun 19, 2024
Accepted: Oct 13, 2024
Available online: Nov 5, 2024
Published: Dec 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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5,141 | https://bio-protocol.org/en/bpdetail?id=5141&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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Peer-reviewed
CRISPR/Cas9-Based Protocol for Precise Genome Editing in Induced Pluripotent Stem Cells
AS Avinash Singh
SB Swathy Babu
MP Marcus Phan
SY Shauna H. Yuan
Published: Vol 14, Iss 24, Dec 20, 2024
DOI: 10.21769/BioProtoc.5141 Views: 778
Reviewed by: Samantha HallerPhilipp WörsdörferRakesh Bam
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Cited by
Original Research Article:
The authors used this protocol in Scientific Reports Apr 2024
Abstract
The advent of clustered regularly interspaced short palindromic repeats (CRISPR)/Cas9-based genome editing has marked a significant advancement in genetic engineering technology. However, the editing of induced pluripotent stem cells (iPSCs) with CRISPR presents notable challenges in ensuring cell survival and achieving high editing efficiency. These challenges become even more complex when considering the specific target site. P53 activation as a result of traditional CRISPR editing can lead to apoptosis, potentially worsening cell health or even resulting in cell death. Mitigating this apoptotic response can enhance cell survival post-CRISPR editing, which will ultimately increase editing efficiency. In our study, we observed that combining p53 inhibition with pro-survival small molecules yields a homologous recombination rate of over 90% when using CRISPR in human iPSCs. This protocol significantly streamlines the editing process and reduces the time and resources necessary for creating isogenic lines.
Key features
• The combination of p53 inhibition and pro-survival small molecules promotes cell survival and increases the efficiency of genome editing.
• Genome editing can be completed in as little as 8 weeks for iPSCs, significantly reducing the total time required.
• Achieves a homologous recombination rate of over 90% in human iPSCs.
Keywords: iPSCs CRISPR Cas9 Nucleofection Editing efficiency Homologous recombination rate
Graphical overview
Outline of Protocol
Background
Genome editing has revolutionized biotechnology by allowing scientists to make precise changes to an organism's DNA. The ability to modify an organism's genome allows experts to study difficult evolutionary and medical problems in more complex systems [1]. The application of gene editing in induced pluripotent stem cells (iPSCs) to create genetically modified isogenic lines can enhance our understanding of how genetic mutations cause disease. Gene editing ranges from conceptually straightforward gene knockouts to more intricate modifications such as point mutations or whole gene insertions. However, currently, gene editing is still confronted with challenges [2–4]. In plant and animal models, clustered interspaced short palindromic repeats (CRISPR) have emerged as the tool of choice for genome editing [5]. Nowadays, the nuclease versions Cas9 and Cas12a are commonly used [1]. The CRISPR-Cas9 and Cas12a systems work with a guide RNA (gRNA) that is divided into two parts: a trans-activating CRISPR RNA (tracrRNA) that binds directly to the Cas nuclease to create ribonucleoprotein (RNP) and a CRISPR RNA (crRNA) that indicates the genomic target location [6]. When compared with previous editing techniques, the CRISPR-Cas innovation offers more precise targeting and editing proficiency as well as being easier and less expensive to configure, allowing it to be used more extensively [7–9]. However, the CRISPR-Cas editing systems still require improvement, particularly for point mutations. In certain cases, there may be several risk variants associated with a target gene, necessitating the creation of multiple lines with distinct single nucleotide polymorphisms (SNPs) or the sequential insertion of genetic modifications. Therefore, to enable more laboratories to produce the needed genetically modified lines, it is essential to develop a gene editing technique that is both highly efficient and easily adaptable. Currently, our attention is on overcoming the barriers to introducing point mutation in cell lines and improving the method to be as effective as possible. Editing efficiency may be increased by preventing cell death with the use of a Rho-related protein kinase (ROCK) inhibitor and blocking the p53 pathway [7]. In this protocol, we used pro-survival small molecules in conjunction with p53 inhibition to attain a homologous recombination rate exceeding 90%.
Materials and reagents
Biological materials
iPSC line (human NDC1) [10]
Reagents
mTeSR Plus basal (Stem Cell Technologies, catalog number: 100-0274)
mTeSR Plus 5× supplement (Stem Cell Technologies, catalog number: 100-0275)
CloneR (Stem Cell Technologies, catalog number: 05888 10 mL)
Revitacell supplement (Gibco, catalog number: A2644501 100×)
1× D-PBS (Gibco, catalog number: 10010049 500 mL)
DMEM/F12 (Gibco, catalog number: A4192001 500 mL)
Matrigel (Corning, catalog number: 47743-706 5 mL)
Accutase (VWR, catalog number: AT104 100 mL)
ReLeSR (Stem Cell Technologies, catalog number: 100-0484)
Alt-R Cas9 HDR enhancer (stock solution 3 mM) (IDT, catalog number: 1081062)
pmaxGFP (LONZA, catalog number: V4XP3032 50 μL at 1 μg/μL)
P3 primary cell nucleofector solution (LONZA, catalog number: V4XP3032 675 μL)
P3 primary cell nucleofector supplement (LONZA, catalog number: V4XP3032 150 μL)
Single-guide RNA (sgRNA) (IDT custom-made, 100 μM)
Alt-R S.p. HiFi Cas9 Nuclease V3 (IDT, catalog number: 10810559 500 μg at 10 μg/μL)
Single-strand oligo donor (ssODN) (IDT, 10 nmol)
shp53-f2 plasmid (Addgene)
Alt-R Cas9 electroporation enhancer (IDT, catalog number: 1075915 100 μm)
Molecular-grade H2O (Cytiva, catalog number: 82007-334 500 mL)
Zymo quick DNATM MicroPrep kit (Zymo Research, catalog number: D3021)
Agarose (VWR Life Sciences, catalog number: N605-500G)
Zymo CleanTM Gel DNA Recovery kit (Zymo Research, catalog number: D4002)
Solutions
Ribonucleoprotein (RNP) complex (see Recipes)
CloneR media (see Recipes)
Nucleofection reaction solution (see Recipes)
Nucleofection media (see Recipes)
P3 nucleofection solution mix (see Recipes)
mTeSR complete media (see Recipes)
Matrigel (see Recipes)
Recipes
RNP complex
This should be freshly prepared.
Reagent Final concentration Volume
Cas9 nuclease V3 64 mM 0.8 μL
sgRNA 100 µM 1.2 μL
D-PBS (1×) 3.0 μL
Total 5.0 μL
CloneR media
This should be freshly prepared. Can be stored at 4 °C for one week.
Media component Volume
mTeSR complete 8.9 mL
Clone R 1 mL
Revitacell 100 µL
Total 10 mL
Nucleofection reaction solution
This should be freshly prepared.
Component Test reaction GFP control No pulse control
P3 primary cell nucleofector solution 1 μL 7 μL 1 μL
shp53-f2 plasmid 1 μL 1 μL 1 μL
Alt-R electroporation enhancer (100 μM) 1 μL 1 μL 1 μL
ssODN (single-stranded oligodeoxynucleotide) (100 μM) 1 μL 0 μL 1 μL
RNP complex 5 μL 0 μL 5 μL
GFP (0.5 μg/μL) 0 μL 1 μL 0 μL
Cells (1 million/reaction) (μL can be altered) 11 μL 11 μL 11 μL
Total 20 μL 20 μL 20 μL
Nucleofection media
This should be freshly prepared.
Component Volume
CloneR media 1 mL
Alt-R Cas9 HDR enhancer 10 µL
P3 nucleofection solution mix
This should be freshly prepared.
Component Volume
P3 primary cell nucleofector solution 16.4 μL
P3 primary cell nucleofector supplement 3.6 μL
Total 20 μL
mTeSR Plus complete media
This should be freshly prepared and can be stored at 4 °C for two weeks.
Component Volume
mTeSR Plus basal 40 mL
mTeSR Plus 5× supplement 10 mL
Total 50 mL
Matrigel
Dilute Matrigel right before the experiment with DMEM/F12. The diluted Matrigel can be stored at 4 °C for 2 weeks.
Component Volume
DMEM F12 11 mL
Matrigel (1 mg/mL) 1 mL
Total 12 mL
Laboratory supplies
6-well culture plate (Thermo Scientific, catalog number: 12556004)
24-well plate (Thermo Scientific, catalog number: 12556006)
12-well plate (Thermo Scientific, catalog number: 12556005)
1.7 mL microtube (Genesee Scientific, catalog number: 24-282C)
1,250 μL tips (GeneMate, catalog number: P-1237-1250)
200 μL tips (Neptune, catalog number: 89140-902)
20 μL tips (GeneMate, catalog number: P-1237-20)
10 μL tips (GeneMate, catalog number: P-1234-10)
15 mL Falcon tubes (Sarstedt, catalog number: 62.554.205)
16-well Nucleocuvette Strip (Lonza, catalog number: V4XP3032)
Equipment
Nucleofector (Lonza, model: 4D)
Centrifuge (Sorvall, model: ST16R)
Gel imager (Bio-Rad, model: ChemiDocTM MP Imaging System)
Gel electrophoresis machine (Labnet, model: Enduro GelXL)
NanoDrop (Thermo Scientific, model: Nanodrop One)
PCR thermocycler (Bio-Rad, model: T100TM thermal cycler)
Microscope (EVOS, Advance Microscopy group)
Incubator (SANYO CO2 incubator)
Biosafety cabinet (Nuaire, model: LabGard, Class II, Type A2, Biological Safety Cabinet)
Hemocytometer (Hausser Scientific Horsham, model: Bright-Line)
Software and datasets
Synthego.com (https://design.synthego.com/#/)
IDT (https://www.idtdna.com/site/order/designtool/index/CRISPR_CUSTOM)
ImageJ (https://ij.imjoy.io/)
Procedure
In silico design sgRNA and ssODN
Design sgRNA and ssODN by using IDT software: https://www.idtdna.com/site/order/designtool/index/HDRDESIGN.
The exact amount to resuspend sgRNA (Table 1) and ssODN (Table 2) to a 100 μM concentration can be determined using IDT Resuspension Calculator: https://www.idtdna.com/calc/resuspension/.
Table 1. Information of guide RNA used in this protocol
dsSNP ID Sequences PAM Site
rs121918393 GAGGCGCACCCGCAGCTCCT CGG
Table 2. Information on HDR templates used in this study
dsSNP ID HDR Template Sequences
rs121918393 GGAGCCGCTTACGCAGCTTGCGCAGGTGGGAGGCGAGGCTCACGCGCAGCTCCTCTGTGCTCTGGCCGAGCATGGCCTGCACCTCGCCGCGGTACTGCAC
Cell culture and gene editing
Maintain the iPSC line in mTeSR Plus complete media (see Recipe 6).
Note: Maintain the iPSC line in mTeSR Plus complete media at 37 °C in a humidified 5% CO2 incubator. Change media every other day. Passage cells when 75%–80% confluence is achieved.
Change cell culture media to mTeSR Plus complete media with additional 1% Revitacell supplement 1 h before the start of nucleofection.
Prepare a Matrigel-coated 24-well plate (see Recipe 7) for the nucleofected cells >1 h prior.
After aspirating Matrigel, add 500 μL of nucleofection media (see Recipe 4). Use half these amounts if using a 48-well plate.
Aspirate media from the iPSC cell culture.
Add 2 mL of 1× D-PBS to clean/rinse the cell monolayer and aspirate.
Add 0.5 mL of Accutase, incubate at 37 °C for <10 min (8 min is optimal), and check that cells have been released using a microscope.
Add 2 mL of DMEM/F12 to neutralize the Accutase.
Gently wash cells off the plate using a p1000 pipette and transfer to a 15 mL conical tube.
Add additional DMEM/F12 to make up to 10 mL.
Take 10 μL of cell suspension and count using a hemocytometer or other cell counting methods. A cell concentration of 1 million cells per reaction is recommended (see cell counting protocol in File S1).
Note: We recommend the user to optimize the cell density when a new line is used.
Take the total number of cells (per milliliter of cell suspension) needed for the experiment depending on how many separate reactions are being done.
Note: For example, three reactions would need 3 million cells in total. If the cell concentration was 0.5 million/mL, then 6 mL would need to be taken from the counted suspension to have 3 million cells in total.
Centrifuge the cell suspension at 200× g for 7 min at 4 °C with acceleration at 9 and deceleration at 5.
Aspirate and resuspend cells in 11 μL per reaction of P3 primary cell nucleofector solution mix (see Recipe 5).
In each Nucleofection reaction mixture, add the cell suspension and 5 μL of the pre-prepared RNP complex and pmaxGFP, ssODN, shp53-f2 plasmid, and Alt-R Cas9 electroporation enhancer and add additional P3 primary cell nucleofector solution if required (see Recipe 3).
Pipette suspension up and down 2–3 times to mix and transfer 20 μL of the nucleofection reaction mixture to the corresponding 16-well nucleocuvette strip wells.
Gently tap the 16-well nucleocuvette strip on the counter inside the biosafety cabinet to ensure that there are no air bubbles.
Turn on the Lonza 4D nucleofector, select X, and select the 16-strip Nucleocuvette with the P3 program CA137.
Note: We recommend optimizing by testing 4–8 programs initially when using a new cell line.
Place the nucleocuvette strip into the Lonza 4D system and nucleofect.
Check corresponding wells to program. Ensure that the no-pulse control does not have an assigned program, then select START. The nucleofection will take only ~30 s (Figure 1). Any errors will show on the screen.
Figure 1. Pictorial presentation of setting Lonza nucleofector for gene editing. Step 1: Set the Lonza nucleofector; select pulse code, solution, and cell type. Step 2: Select the program. Step 3: Run the program after selecting all required wells. Step 4: Completion of nucleofection.
Remove the Nucleocuvette from the nucleofector and incubate at 37 °C for 10–20 min.
Notes:
Do not add anything, move, or disturb the cuvette strip during this time.
This allows the lipid bilayer of the cells to reposition/organize.
After 10–20 min incubation, add 20 μL of the prewarmed nucleofection media (see Recipe 4) from the pre-prepared 24-well plate to each well in the nucleocuvette.
Using a p20 pipette, add the 40 μL nucleoporated cell suspension into the 500 μL of nucleofection media already in the Matrigel-coated 24-well plate.
Incubate cells at 37 °C; then, check and change media at 24 h with CloneR media (see Recipe 2).
Forty-eight hours after nucleofection, replace the CloneR media with fresh CloneR media. A complete change is not required. This is to compensate for the evaporation/degradation of previous CloneR.
Post-nucleofection
Twenty-four hours after nucleofection, count the percentage of GFP-expressing cells at 10× magnification. Take two images (Figure 2) with the brightfield filter and two images of the same field of view with the GFP filter per reaction (well). Repeat in 48 h. Save these images onto a USB to count using ImageJ software. After 48 h, the ability to see GFP under the microscope filter slowly dissipates.
Figure 2. Induced pluripotent stem cell (iPSC) colonies with pmaxGFP 24 h post-nucleofection GFP vs. brightfield at 10× magnification
Once cells reach 70%–80% confluency, extract genomic DNA using the Zymo quick DNATM MicroPrep kit.
Quantify genomic DNA levels using a Nanodrop and perform PCR for the desired gene.
Note: For achieving high-quality results in genomic DNA applications like PCR, it is crucial to use good-quality DNA and ensure the following practices: quantification of genomic DNA, optimization of PCR conditions, and running a gel for verification.
Run 20 μL of the PCR product on 1% agarose gel to check the quality of the PCR product.
Cut the PCR product and purify by Zymo CleanTM Gel DNA Recovery kit.
Quantify the eluate using Nanodrop to determine the optimal amount for Sanger sequencing.
Results from Sanger sequencing can be analyzed using Synthego’s ICE online software and/or gene viewer programs, e.g., SnapGene.
To analyze using ICE, download the ab.1 file for the desired sample to analyze and the corresponding control file.
Go to https://ice.synthego.com and drag and drop the ab.1 files into the corresponding control and experiment sample boxes.
Use the guide and repair ssODN sequences.
Select add samples to analysis and analyze (Figure 3).
Figure 3. Sanger sequence view showing edited and control sequences in the region around the guide sequence. This shows sequence base calls from both the control and the edited sample .ab1 files. The horizontal black underlined region represents the guide sequence. The horizontal red underline is the PAM site. The vertical black dotted line represents the actual cut site.
After confirmation of gene editing, cells can be dissociated and isolated into single cells.
Single clone selection
Coat a 10 cm plate with Matrigel for <1 h.
Aspirate media from bulk cell plate, wash with 2 mL of 1× D-PBS, and aspirate.
Add 0.5 mL (for a 6-well plate) and 0.125 mL (for a 24-well plate) of Accutase for <10 min at 37 °C.
Add 1 mL of DMEM/F12 to the well to wash and separate.
Transfer cell suspension to a 15 mL conical tube and top up to 10 mL with DMEM/F12.
Note: This further dilutes the dissociation reagent to ensure cell health.
Centrifuge at 100× g for 5 min.
Aspirate supernatant and flick tube to loosen cell pellet.
Add 1 mL of cloning media to the cell pellet and pipette to mix.
Count the cells by hemocytometer (File S1).
Seed two 10 cm cell culture dishes with 100 and 300 cells in 10 mL of media.
Keep the plate in the incubator at 37 °C for 48–72 h.
Examine the cells under the microscope after 48 h to see if they are single cells or not.
Cells should be ready for picking after 72–96 h of seeding (when cells form single colonies) (Figure S1).
After 72–96 h, when small colonies originating from single cells appear, prepare the microscope in the biosafety cabinet.
Prepare a 24-well plate with Matrigel media <1 h prior to picking.
With the appropriate plate under the microscope (Figure 4) at 4× magnification, use a p200 pipette on the 100 μL setting (to pick a 24-well), and pick colonies that are clearly isolated with a good size (up to 5–10 μm) and shape (Video S1).
Notes:
Cell colony may need to be lightly scraped to lift it off the plate before pipetting up.
After several colonies have been picked, the media in the plate wells may need to be topped up in order to continue picking.
Figure 4. Image of microscope setup inside the biosafety cabinet for colony pickup
Allow colonies in each well to form large, good sized colonies (this can take 3–5 days post-picking).
Once the colonies are big enough to be split, prepare/prewarm a new Matrigel-coated 24-well plate with 500 μL of cloning media in each well.
Split corresponding cells into a 12-well plate from a 24-well plate by using ReLeSR as a digestive agent and extract genomic DNA from half of the cells.
After genomic DNA extraction, perform a standard PCR, send the sample for sequencing, and verify with a sequencing analysis program or perform ICE analysis (the same as steps 7 to 11).
Note: We use commercially available genomic DNA extraction kits for better DNA quality.
After confirmation, perform downstream experiments from the edited cell lines.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article:
Singh et al. [11]. A high efficiency precision genome editing method with CRISPR in iPSCs. Scientific Reports.
General notes and troubleshooting
General notes
A passage number below p25 will result in better editing efficiency.
On the day of nucleofection, the media should be changed to mTeSR plus with (1%) Revitacell supplement.
Try to use the IDT resuspension calculator to resuspend the sgRNA and ssODN.
Before use, these reagents should be centrifuged for a brief amount of time.
The Cas9 enzyme should always be taken out from the -20 °C freezer directly before use such as when making the RNP complex.
If using Accutase to detach, you should use double the volume of DMEM/F12 media to dilute the Accutase.
After nucleofection, immediately keep the vessel in the incubator at 37 °C for 10–20 min.
For single-cell colony picking, seed 100 and 300 cells in two different 10 cm plates. For single-cell colony picking, seeding cells at the correct density is crucial to ensure that individual cells are spaced far enough apart to grow into isolated colonies. Our approach to seed 100 and 300 cells in two different 10 cm plates is a strategy to balance cell density and the ease of picking single colonies.
It is highly recommended to cut the desired PCR product from the gel, purify it, and send it for sequencing.
Troubleshooting
Problem 1: Low cell survival rate after nucleofection.
Possible cause: i) The cell number is low.
Solution: i) Use the appropriate cell number with a minimum of 1 million cells per reaction.
Possible cause: ii) Issue with Cas9 concentration.
Solution: ii) We recommend optimizing the Cas9 concentration by using different concentrations of Cas9 in a titration experiment.
Possible cause: iii) Harsh pipetting and centrifugation.
Solution: iii) Pipette very gently during the cell handling and perform centrifugation with centrifugation breaking methods.
Problem 2: Low editing efficiency.
Possible cause: The guide RNA or ssODN may not be working. Cas9 concentration may be incorrect.
Solution: i) Always try 2–3 different guideRNAs and ssODN.
Solution: ii) Optimize Cas9 concentration.
Problem 3: Poor sequencing results from Sanger sequencing.
Possible cause: Problems with PCR primers or the selection of the correct PCR product.
Solution: Redesign the PCR primer and always run an agarose gel, cut the expected size fragment, elute with the gel recovery kit, quantify, and send for sequencing.
Problem 4: Cells are not attaching in the 10 cm dish after plating the single cell suspension.
Possible cause: Lack of cell survival molecules.
Solution: Use CloneR in the growth media.
Acknowledgments
We acknowledge NIH grant 1R03AG070415, Veterans Administration Merit Research Award 1 I01 BX006391, University of Minnesota Faculty Research Development Grant, funds from the University of Minnesota, Institute of Translational Neuroscience (ITN) to SHY. SHY is an ITN Scholar. Figures were created with BioRender.com. We also acknowledge our previous work published in Scientific Reports (2024), DOI: https://doi.org/10.1038/s41598-024-60766-4 on which the current protocol is based [11].
Competing interests
There are no conflicts of interest or competing interests.
References
Robb, G. B. (2019). Genome Editing with CRISPR‐Cas: An Overview. Curr Protoc Essent Lab Tech. 19(1): e36.
Khalil, A. M. (2020). The genome editing revolution: review. J Genet Eng Biotechnol. 18(1): 68.
Carroll, D. (2014). Genome Engineering with Targetable Nucleases. Annu Rev Biochem. 83(1): 409–439.
Urnov, F. D., Miller, J. C., Lee, Y. L., Beausejour, C. M., Rock, J. M., Augustus, S., Jamieson, A. C., Porteus, M. H., Gregory, P. D., Holmes, M. C., et al. (2005). Highly efficient endogenous human gene correction using designed zinc-finger nucleases. Nature. 435(7042): 646–651.
Li, X. L., Li, G. H., Fu, J., Fu, Y. W., Zhang, L., Chen, W., Arakaki, C., Zhang, J. P., Wen, W., Zhao, M., et al. (2018). Highly efficient genome editing via CRISPR–Cas9 in human pluripotent stem cells is achieved by transient BCL-XL overexpression. Nucleic Acids Res. 46(19): 10195–10215.
Budde, J. P., Martinez, R., Hsu, S., Wen, N., Chen, J. A., Coppola, G., Goate, A. M., Cruchaga, C. and Karch, C. M. (2017). Precision genome-editing with CRISPR/Cas9 in human induced pluripotent stem cells. bioRxiv. doi.org/10.1101/187377.
Ihry, R. J., Worringer, K. A., Salick, M. R., Frias, E., Ho, D., Theriault, K., Kommineni, S., Chen, J., Sondey, M., Ye, C., et al. (2018). p53 inhibits CRISPR–Cas9 engineering in human pluripotent stem cells. Nat Med. 24(7): 939–946.
Conti, A. and Di Micco, R. (2018). p53 activation: a checkpoint for precision genome editing? Genome Med. 10(1): 66.
Uffelmann, E., Huang, Q. Q., Munung, N. S. (2021). Genome-wide association studies. Nat Rev Methods Primers. 1: 59.
Israel, M. A., Yuan, S. H., Bardy, C., Reyna, S. M., Mu, Y., Herrera, C., Hefferan, M. P., Van Gorp, S., Nazor, K. L., Boscolo, F. S., et al. (2012). Probing sporadic and familial Alzheimer’s disease using induced pluripotent stem cells. Nature. 482(7384): 216–220.
Singh, A., Smedley, G. D., Rose, J. G., Fredriksen, K., Zhang, Y., Li, L. and Yuan, S. H. (2024). A high efficiency precision genome editing method with CRISPR in iPSCs. Sci Rep. 14(1): 9933.
Supplementary information
The following supporting information can be downloaded here:
File S1. Protocol for cell counting by using hemocytometer slide
Figure S1. Image of single cell
Video S1. Video of colony piking under the microscope
Article Information
Publication history
Received: Aug 20, 2024
Accepted: Oct 14, 2024
Available online: Nov 5, 2024
Published: Dec 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Peer-reviewed
Live Visualization of Calcified Bones in Zebrafish and Medaka Larvae and Juveniles Using Calcein and Alizarin Red S
RK Rina Koita
SO Sae Oikawa
TT Taisei Tani
MM Masaru Matsuda
AK Akinori Kawamura §
(§ Technical contact)
Published: Vol 14, Iss 24, Dec 20, 2024
DOI: 10.21769/BioProtoc.5142 Views: 513
Reviewed by: Alberto RissoneAmr Galal Abdelraheem Ibrahim Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Proceedings of the National Academy of Sciences of the United States of America Jun 2024
Abstract
Zebrafish and medaka are valuable model vertebrates for genetic studies. The advent of CRISPR-Cas9 technology has greatly enhanced our capability to produce specific gene mutants in zebrafish and medaka. Analyzing the phenotypes of these mutants is essential for elucidating gene function, though such analyses often yield unexpected results. Consequently, providing researchers with accessible and cost-effective phenotype analysis methods is crucial. A prevalent technique for investigating calcified bone development in these species involves using transgenic fish that express fluorescent proteins labeling calcified bones; however, acquiring these fish and isolating appropriate crosses can be time-consuming. We present a comprehensive protocol for visualizing ossified bones in zebrafish and medaka larvae and juveniles using calcein and alizarin red S staining, which is both economical and efficient. This method, applicable to live specimens during the ossification of bones, avoids apparent alterations in skeletal morphology and allows for the use of different fluorescent dyes in conjunction with transgenic labeling, thus enhancing the analysis of developmental processes in calcifying bones, such as vertebrae and fin rays.
Key features
• The calcified bones of alive zebrafish and medaka larvae and juveniles can be visualized repeatedly using simple and inexpensive calcein and alizarin red S.
• No need to use transgenic fish to visualize ossified bones, allowing for rapid analysis of bone phenotypes in mutants.
• Double staining is possible in transgenic fish with reporter genes such as GFP and DsRed using alizarin red S or calcein, which exhibit different fluorescence.
• Ossification processes of bones such as vertebrae, ribs, and fin rays can be analyzed in mutants.
Keywords: Zebrafish Medaka Calcified bones Live staining Calcein Alizarin red S
Graphical overview
Background
Zebrafish and medaka are model vertebrates that are amenable to genetic approaches. Chemical mutagenesis has led to the isolation of hundreds of mutants in these species [1,2], contributing greatly to our knowledge of vertebrate development and other biological phenomena. The development of CRISPR-Cas9 technology has further enhanced our ability to generate mutants of specific genes in vertebrate models [3–5]. To fully understand the function of a gene, it is crucial to analyze the phenotype of the mutants. However, the generation of mutants often results in unexpected phenotypes. Therefore, it is important to provide researchers with accessible and cost-effective methods to analyze the phenotypes.
A common approach to studying the development of calcified bones in zebrafish and medaka is to use transgenic fish that specifically express fluorescent proteins labeling calcified bones [6,7]. However, it is necessary to first obtain transgenic fish, especially for researchers who do not specialize in bone research. Even if they manage to obtain them, it takes several months to analyze them because new fish have to be isolated by crossing transgenic fish with the mutant fish. Here, we describe a detailed step-by-step protocol for visualizing the ossified bones of zebrafish and medaka larvae and juveniles. This method allows the visualization of calcified bones in zebrafish and medaka by staining with calcein and alizarin red S, which are inexpensive reagents. The staining can be performed in one or two days on live zebrafish and medaka and can be repeated multiple times, as it does not cause any noticeable abnormalities in the subsequent skeletal morphology such as vertebrae and fin rays [8,9]. In addition, calcein and alizarin red S are different fluorescent dyes, and double staining is possible by using different fluorescent dyes depending on the fluorescent protein in transgenic fish. Analysis of mutants using this method is expected to improve our understanding of various developmental processes in the calcifying bones of zebrafish and medaka, including vertebrae and fin rays.
Materials and reagents
Biological materials
Zebrafish (Danio rerio, Riken Wild-type (RW) strain, the National BioResource Project Zebrafish in Japan)
Medaka (Oryzias latipes, Hd-rR strain, the National BioResource Project Medaka in Japan)
Reagents
Calcein (Fujifilm Wako, catalog number: 340-00433)
NaOH (Fujifilm Wako, catalog number: 194-18865)
NaCl (Fujifilm Wako, catalog number: 191-01665)
KCl (Fujifilm Wako, catalog number: 160-03555)
CaCl2 (Fujifilm Wako, catalog number: 038-19735)
HEPES (Fujifilm Wako, catalog number: 342-01375)
Alizarin red S (Fujifilm Wako, catalog number: 011-01192)
KOH (Fujifilm Wako, catalog number: 168-21815)
Tricaine (Fujifilm Wako, catalog number: 051-06571)
2-Phenoxyethanol (Fujifilm Wako, catalog number: 163-12075)
Methylcellulose 1500 cP (Fujifilm Wako, catalog number: 139-02145)
Solutions
2.0% calcein stock solution (see Recipes)
0.2% calcein staining solution (see Recipes)
1× Ringer’s solution (see Recipes)
1/3× Ringer’s solution (see Recipes)
0.1% alizarin red S stock solution (see Recipes)
0.01% alizarin red S staining solution (see Recipes)
Tricaine (MS-222) anesthetizing stock solution for zebrafish (see Recipes)
2.0% methylcellulose solution (see Recipes)
Recipes
2.0% calcein stock solution
Reagent Final concentration Amount
Calcein 2.0% 1 g
0.5 N NaOH solution Adjust to pH 7.0
1/3 Ringer’s solution (Recipe 4) n/a Add up to 50 mL
Total n/a 50 mL
The prepared 2.0% calcein stock solution should be aliquoted into small tubes and stored at -20 °C in the dark. Instead of 1/3 Ringer’s solution, E3 or similar buffers can be used. Prepare a 0.5 N NaOH solution by dissolving 1.0 g of NaOH in 50 mL of distilled water.
0.2% calcein staining solution
Reagent Final concentration Amount
2.0% calcein stock solution (Recipe 1) 0.2% 300 μL
1/3 Ringer’s solution (Recipe 4) n/a 2.7 mL
Total n/a 3.0 mL
0.2% calcein staining solution should be prepared prior to staining. The example shown above is for a 35 mm dish; the volume to be added is 3.0 mL. If more dishes are used, the volume can be increased. Instead of 1/3 Ringer’s solution, E3 or similar buffers can be used.
1× Ringer’s solution
Reagent Final concentration Amount
NaCl 116 mM 6.78 g
KCl 2.9 mM 0.216 g
CaCl2·2H2O 1.8 mM 0.265 g
HEPES 5.0 mM 1.192 g
0.5 N NaOH solution Adjust to pH 7.0
DW n/a Add up to 1,000 mL
Total n/a 1,000 mL
1× Ringer’s solution is stable at room temperature for years.
1/3× Ringer’s solution
Reagent Final concentration Amount
1× Ringer’s solution (Recipe 3) 1/3× 333 mL
DW n/a Add up to 1,000 mL
Total n/a 1,000 mL
1/3× Ringer’s solution is stable at room temperature for years.
0.1% alizarin red S stock solution
Reagent Final concentration Amount
Alizarin red S 0.1% 0.05 g
1 N KOH solution Adjust to pH 7.4
1/3 Ringer’s solution (Recipe 4) n/a Add up to 50 mL
Total n/a 50 mL
0.1% alizarin red S stock solution should be stored at room temperature in the dark. Instead of 1/3 Ringer’s solution, E3 or similar buffers can be used. Prepare a 1 N KOH solution by dissolving 2.0 g of KOH in 50 mL of distilled water.
0.01% alizarin red S staining solution
Reagent Final concentration Amount
0.1% alizarin red S stock solution (Recipe 5) 0.01% 300 μL
1/3 Ringer’s solution (Recipe 4), or DW n/a 2.7 mL
Total n/a 3.0 mL
0.01% alizarin red S staining solution should be prepared prior to staining. The example shown above is for a 35 mm dish; the volume to be added is 3.0 mL. If more dishes are used, the volume can be increased. Instead of 1/3 Ringer’s solution, E3 or similar buffers can be used.
Tricaine (MS-222) anesthetizing stock solution for zebrafish
Reagent Final concentration Amount
Tricaine 2.0% 1 g
DW Add up to 50 mL
Total n/a 50 mL
The prepared tricaine anesthetizing stock solution should be divided into small tubes and stored at -20 °C in the dark.
2.0% Methylcellulose solution
Reagent Final concentration Amount
Methylcellulose 1500 cP 2.0% 1 g
DW n/a Add up to 50 mL
Total n/a 50 mL
To completely dissolve the methylcellulose in DW, shake vigorously for several days. When photographing, the fish is placed in this viscous solution to orient the fish in the desired position. It is possible to adjust the solution to a percentage of 2%–3% methylcellulose depending on personal preference.
Laboratory supplies
Non-treated dish, 35 mm (IWAKI, catalog number: 1000-035)
Glass-base dish, 35 mm (IWAKI, catalog number: 3970-035)
Transfer pipette, 3 mL (BD Falcon, catalog number: 357575)
Equipment
Fluorescent stereomicroscope (Leica, model: M205 FA)
Digital camera (Leica, model: DFC350 FX)
Software and datasets
ImageJ (v. 1.53e, September 2020, free to use, https://imagej.net/ij/)
Procedure
Both calcein (green fluorescent signal) and alizarin red S (red fluorescent signal) can be used to visualize calcified bones in live zebrafish and medaka. However, calcein is recommended, except in cases such as staining the bones of transgenic fish with the GFP gene. Calcein has a shorter staining time, higher signal-to-noise ratio, and stronger fluorescent intensity than alizarin red S. Visualization of the bones in the internal body, such as vertebrae, is possible until the scales have calcified. In the following protocol, staining methods are essentially the same for zebrafish and medaka, unless otherwise noted. When referring to “fish,” both zebrafish and medaka are included in this protocol.
Calcein staining in live zebrafish and medaka larvae and juveniles
Obtain fertilized embryos by crossing female and male adult fish.
Raise the fish embryos to the desired developmental stages for analysis. For more information on the developmental stages of zebrafish and medaka, see [10–12].
Use a 35 mm dish for calcein staining. Each dish can accommodate up to five fish for staining. To ensure an even staining, avoid placing more than six fish in one dish. Prepare the necessary number of 35 mm plastic dishes.
Prepare 3 mL of 0.2% Calcein staining solution per dish to be analyzed. Adjust the volume of the staining solution based on the number of dishes. After preparation, store the solution at room temperature in the dark.
In addition, make sure to have plenty of room-temperature water (possibly with 1/3 Ringer’s solution or tap water that has been left for at least one day to remove sodium hypochlorite) for washing purposes.
First, place the fish whose bones need to be visualized in a 35 mm dish filled with enough water. When transferring the fish, use a 3 mL transfer pipette with the tip removed, allowing the larvae to pass through without damage. Carefully transfer the fish to the 35 mm dish.
Next, remove water using a regular 3 mL pipette (uncut tip). Shift the lid slightly to prevent the fish from jumping while removing the water.
Immediately add 3 mL of 0.2% Calcein staining solution to the 35 mm dish and stain the fish at room temperature for 15 min. Cover the dish with a lid to prevent the fish from jumping during staining (Figure 1A).
Figure 1. Staining of calcified bones in live juvenile zebrafish. To stain the ossified bones, a live juvenile zebrafish is placed in a 35 mm dish and stained with calcein (A) or alizarin red S (B). For transferring the fish, a 3 mL transfer pipette with the tip cut off is used, while a separate 3 mL transfer pipette with the tip intact is used to transfer the solution.
Discard the 0.2% Calcein staining solution using a 3 mL transfer pipette. Immediately add approximately 4 mL of water to the 35 mm dish and mix gently.
Remove the water using a 3 mL transfer pipette. Immediately add another 4 mL of water into the 35 mm dish and mix gently. Place the fish in the 35 mm dish for 5 min. Repeat this process three times until the yellow dye of the calcein disappears.
Keep the stained fish in water in the 35 mm dishes. Fluorescent images can be captured immediately after staining. However, it is recommended that pictures be taken the day after staining to reduce background signals and obtain high signal-to-noise ratio images. When photographing the next day, place the fish in an environment with the optimum temperature and do not feed them until the photographs have been taken.
Stained calcein signals can still be detected in areas that were calcified at the time of the initial procedure, even after 3 days. The pulse-chase experiments utilizing dual labeling with alizarin red S allow for the analysis of the ossification process in the bones of interest. For details, see Akama et al. [9].
Alizarin red S staining of zebrafish and medaka larvae and juveniles
The alizarin red S staining method is basically the same as calcein staining, except for the staining time.
Perform the same procedures as described in steps A1–7.
Immediately add 3 mL of 0.01% alizarin red S staining solution to the 35 mm dish and stain the fish at room temperature for 60 min. Cover the dish with a lid to prevent the fish from jumping during staining (Figure 1B).
Perform the same procedures as described in steps A9–11.
Anesthesia treatment of zebrafish and medaka prior to imaging
Administering anesthesia to zebrafish and medaka larvae and juveniles is a critical and potentially risky step in this protocol, so caution must be exercised to avoid overdosing the fish. It is recommended to administer anesthesia to one fish at a time just before capturing fluorescent images. The amount of anesthesia required depends on the size of the fish, so it is advisable to anesthetize the smaller fish first.
For zebrafish, prepare a new 35 mm dish by adding 3 mL of water and 10 μL of tricaine in the room with the fluorescent stereomicroscope. For medaka, prepare a new 35 mm dish by adding 3 mL of water and 3 μL of 2-phenoxyethanol.
Prepare the stained fish in 35 mm dishes.
For fluorescence imaging, prepare a 35 mm glass-base dish filled with a 2% methylcellulose solution in the glass section.
Using a 3 mL transfer pipette with the tip removed, carefully transfer one fish to the 35 mm dish with the anesthesia.
Ensure that the fish are immobilized by the anesthesia treatment. If the fish are still moving, add more anesthesia reagents until they are still. Again, it is important to be cautious with the amount of anesthesia given, as excessive amounts of anesthesia can be fatal for larvae and juveniles. The anesthetic solution can be used for other fish repeatedly.
Using a 3 mL transfer pipette with the tip removed, carefully transfer one fish to a 35 mm glass base dish filled with 2% methylcellulose solution for imaging.
Imaging of stained ossified bones of zebrafish and medaka using a fluorescent stereomicroscope
Zebrafish and medaka grow to over 5 mm in length during the ossification of their skeletons. Fluorescent images of stained calcified bones can be observed using a fluorescent stereomicroscope. It is important to note that calcein or alizarin red S staining does not cause obvious developmental abnormalities in zebrafish and medaka, so the fish can be recovered and grown normally after the photographs are taken. Additionally, calcein signals can be captured under the same conditions as normal GFP signals, while alizarin red S signals can be captured under the same conditions as normal DsRed signals.
When transferring anesthetized fish to a 2% methylcellulose solution in the 35 mm dish, it is important to be aware that the anesthesia wears off after several minutes, causing the fish to start moving. Therefore, it is crucial to efficiently capture the desired fluorescent images within a limited time. If the fish begin to move during the analysis, it is advisable to re-anesthetize them.
The standard length (SL), which is the length of the fish from the tip of the snout to the posteriormost region of the body where caudal fin rays insert, is typically used to indicate the developmental stages of larvae and juvenile fish [12]. Thus, it is crucial to start by capturing a clear image of the entire stained fish. Position the anesthetized fish, include a scale bar for reference, and take a lateral image of the entire fish.
Capture the desired fluorescent images by adjusting the orientation of the stained fish and changing the magnification (Figures 2, 3).
Figure 2. Visualization of the calcified bones in live juvenile zebrafish. (A–E) Calcein staining was performed for live zebrafish juveniles. (F) Alizarin red S staining. All the images are lateral views. Developmental stages of the stained fish are standard length (SL) 7.5 mm (A), SL 7.7 mm (B), SL 8.0 mm (C), SL 10.5 mm (D), SL 8.0 mm (E), and SL 7.8 mm (F). Scale bars: 500 μm.
Figure 3. Visualization of the calcified bones in live medaka juveniles. (A–C) Calcein staining was performed for live medaka juveniles. (D) Alizarin red S staining. Developmental stages of the stained fish shown in (A–D) are stage 41 according to Iwamatsu’s paper [10]. All the images are lateral views. Scale bars: 500 μm.
After finishing the photography, return the fish to the original 35 mm dish using a 3 mL transfer pipette without the tip. To supply oxygen to the fish and remove the viscous methylcellulose from its body, carefully apply water from the 35 mm dish onto the fish’s mouth and gills using a 3 cm transfer pipette. Once you have confirmed that the fish has recovered, cover the dish with a lid and proceed to anesthetize the remaining fish for analysis.
If you wish to continue monitoring the progress, place each fish separately in a 35 mm dish or a 6-well dish and provide them with food and care. You can observe the calcified bone regularly by re-staining using calcein or alizarin red S, with several days between each observation.
The procedures described above can be performed similarly for wild-type zebrafish (Figure 2), medaka (Figure 3), transgenic fish (Figure 4), and mutants (Figure 5).
Figure 4. Example of alizarin red S staining in GFP-expressing transgenic zebrafish. (A–C) Transgenic juvenile zebrafish displaying a strong GFP signal in the proximal regions of the anal fin ray was stained with alizarin red S. The developmental stage of the stained fish is standard length (SL) 7.1 mm. Lateral views. Scale bars: 500 μm.
Figure 5. Examples of calcein staining in zebrafish mutant analysis. (A–D) Calcein staining was performed to distinguish the phenotypes of wild-type and mutant zebrafish. (A, B) Abnormal vertebral morphologies between wild-type and tbx6 mutants. (C, D) The anal fin in hoxc12a;c13a;c12b;c13b mutants is expanded toward the posterior compared with that of the wild type. For details on the mutant phenotypes, see Ban et al. [13] and Adachi et al. [8]. The developmental stages of the stained fish are standard length (SL) 7.5 mm (A, B) and SL 6.9 mm (C, D). Lateral views. Scale bars: 500 μm.
Data analysis
Measurements of the standard length of the stained fish by ImageJ
Use a photograph of a fish taken from the lateral side, showing the entire body, with a scale bar included.
Open the image file in ImageJ software. FIJI, which is a version of ImageJ with additional plugins, can also be used.
Click on the Straight button and measure the length of the scale bar as a reference.
Click on the Analyze menu and select the Set Scale button.
In the Known distance, enter the length of the measured scale bar and click on OK.
Press the Straight button and measure the length of the stained fish from the tip of the snout to the posteriormost region of the body where caudal fin rays are inserted (Figure 6).
Click on the Analyze menu and select Measure; then, the SL is obtained.
Figure 6. Measurement of the standard length (SL) in juvenile zebrafish by ImageJ. The yellow line corresponds to the standard length.
Measurements of the fluorescent signal intensities of the stained fish by ImageJ
Fluorescence intensities can be measured in specific areas of fish stained with calcein or alizarin red S. In this example, we will focus on a fin ray of the anal fin stained with calcein.
Open the image you wish to analyze in ImageJ.
Click on the Image menu and select Color - Split Channels. This will generate three images, each representing one of the RGB channels.
For calcein-stained fish, use the image of the green channel. For fish stained with alizarin red S, use the image of the red channel.
Select the Segmented Line tool and draw lines down the center of the area you wish to analyze (Figure 7A).
Figure 7. Measurement of signal intensities of fish stained with calcein by ImageJ. (A) Example of analyzing the fluorescence intensities of a fin ray using ImageJ. The yellow segmented lines indicate the fin ray to be analyzed (indicated by the arrowhead), extending from the proximal to the distal region. (B) A graph showing the signal intensities along the drawn lines.
Click on the Analyze menu and select Plot Profile. A graph showing the signal intensities along the drawn lines will appear (Figure 7B).
To ensure valid comparisons, use images taken under the same conditions.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article:
Adachi et al. [8]. Teleost Hox code defines regional identities competent for the formation of dorsal and anal fins. Proc Natl Acad Sci USA. (Figure 2, panels C–E, J–L, Figure 3, panels A–C, G, H, N, O, Figure 4, panels H–K).]
General notes and troubleshooting
This protocol outlines a method for visualizing calcified bone in live larvae and juveniles of zebrafish and medaka. For more examples obtained with this method, please refer to our previous papers [8,9,13,14].
When analyzing the whole skeletal structure in adult fish, we recommend using alizarin red and alcian blue staining or an X-ray micro-CT scan [15,16].
Chemical fixation with paraformaldehyde or other agents results in a loss of transparency in these larvae and juveniles, making it difficult to observe the fluorescent signals of their internal skeletons in fixed specimens. Additionally, when larvae and juveniles that have been stained are fixed, their transparency is similarly compromised, rendering them unsuitable for detailed observation of fluorescent signals.
Although pigment cells are observed during the developmental stages of staining, a sufficient fluorescent signal can be detected in their presence. Therefore, it is not necessary to prepare a treatment that inhibits the formation of pigment cells or to use specific strains in which pigment cells do not form.
In later developmental stages, sufficient fluorescent signal can be obtained by staining under the same staining conditions and at the same concentrations described above.
Zebrafish scale calcification begins around SL 8.0 mm, starting in the posterior portion of the body and gradually spreading throughout [17]. In medaka, the ossification of the scales is first observed at stage 42 [10]. Before scale calcification occurs, the internal skeletons, including the vertebrae, can be stained and visualized.
If the fish do not stain adequately, or if the staining process kills the fish, first check whether the reagents you have prepared are correctly made. If sufficient staining is still not achieved, we recommend increasing the staining time.
It is worth noting that stained bones can be observed at a higher resolution using confocal laser microscopy. However, compared to fluorescence stereomicroscopy, imaging with confocal microscopy requires more time and the fish must be kept stationary. Therefore, it is necessary to anesthetize the fish during imaging, and the survival rate of the fish after imaging is also lower than with the fluorescence stereomicroscope.
Although the present protocol is applied to zebrafish and medaka larvae and juveniles, the same method may be adaptable for other fish.
Acknowledgments
We thank the National BioResource Project Zebrafish and the National BioResource Project Medaka for providing fish. This work was supported by KAKENHI Grants-in-Aid for Scientific Research 18K06177, 23K05790 to A.K. and Narishige Zoological Science Award 2021 to A.K. This protocol was adapted and modified from Adachi et al. [8].
Competing interests
The authors declare no competing interests.
Ethical considerations
All experimental procedures using live zebrafish were performed in accordance with the regulations and were approved by the Animal Care and Use Committee of Saitama University. All experiments using medaka were conducted under the guidelines of the Institutional Animal Care and Use Committee of Utsunomiya University.
References
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Article Information
Publication history
Received: Aug 14, 2024
Accepted: Oct 15, 2024
Available online: Nov 4, 2024
Published: Dec 20, 2024
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© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Cryo-SEM Investigation of Chlorella Using Filter Paper as Substrate
PW Peng Wan
MT Meiyue Tao
YZ Yumeng Zhou
WH Wenjun Han
JW Jianxia Wang
JW Jinghan Wang
Published: Vol 14, Iss 24, Dec 20, 2024
DOI: 10.21769/BioProtoc.5143 Views: 230
Reviewed by: Alba BlesaSimab Kanwal Anonymous reviewer(s)
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Abstract
Cryo-electron microscopy (cryo-EM) is a powerful technique capable of investigating samples in a hydrated state, compared to conventional high-vacuum electron microscopy that requires samples to be completely dry. During the drying process, numerous features and details may be lost due to damage caused by dehydration. Cryo-EM circumvents these problems by cryo-fixing the samples, thereby retaining the intact and original features of hydrated samples. This protocol describes a step-by-step cryo-scanning electron microscopy (cryo-SEM) experimental procedure with Chlorella sorokiniana as the subject. By employing filter paper as the sample substrate, we propose a simple and reliable method for cryo-fixation and freeze-fracture of Chlorella sorokiniana in water suspension. The advantage of using filter paper as a substrate lies in its ability to support a thin film of sample, enabling a cold knife to make a cut effortlessly and produce a clean freeze-fractured surface for SEM investigation. By following the approach described in this protocol, both the internal structure and surface morphology of Chlorella sorokiniana can be easily resolved with high quality. This protocol is highly versatile and can be applied to samples dispersed in water or solvents, including cyanobacterial cells, algal cells, and any kind of sample that can be adsorbed onto filter paper.
Key features
• Introducing a reliable way for ideal freeze-fracture of a water-suspended sample using filter paper as substrate.
• Detailed step-by-step descriptions of the entire experiment, covering how to operate the instruments and including some practical experimental tips.
Keywords: Cryo-SEM Algae Chlorella Freeze fracture Filter paper
Graphical overview
Background
Cryo-SEM is an irreplaceable approach for investigating biological samples in a hydrated state [1–5], and it has been proven to be a powerful tool for studying the microstructure and surface morphology of algae and cells [6–11]. There are four main procedures for sample treatment in cryo-SEM, including cryo-fixation, freeze-fracture, freeze-etching, and sputter coating [12,13]. Cryo-fixation is a technique for rapidly freezing a sample to preserve its structure in a near-native state [14,15]. Typically, a liquid nitrogen bath at atmospheric pressure, liquid nitrogen slush, liquid nitrogen under high pressure (for high-pressure freezing), and liquid ethane and propane are employed as cryogenic substances [14–17]. Freeze-fracture is a technique that fractures the cryo-fixed sample in order to reveal its internal structure, typically by a cold knife or a cryo-ultramicrotome [18]. Freeze-etching is the process of getting the fractured surface etched by sublimation of ice, which reveals fine details of the sample's structure. Sputter coating in cryo-state is similar to the conventional sputter coating, except that the sample is maintained in a cryogenic state.
There are some valuable protocols that have been published regarding plant and animal tissues as well as other biomaterials [1–6,9,13]. Nevertheless, only a limited number of protocols are available for cryo-SEM of algae. Research articles regarding cryo-SEM investigation of algae samples usually include a brief description of the cryo-SEM experiments but do not provide much information about the detailed and visualized experimental procedures. Particularly, information such as how the sample is prepared and how it is freeze-fractured is crucial for obtaining fine details of the internal structures.
The technique of freeze-fracture has been widely used in a broad range of samples, such as fluid samples, hydrated polymers and microgels, and plant tissues [18–20]. In the freeze-fracture procedure, a cold knife in a cryogenic state is commonly employed as the cutting tool: a blade with a sharp edge that is positioned above the sample. It sweeps across the top of the sample in parallel with the sample holder. When it hits the sample, preferably a prominent part or a tip, the sample is then freeze-fractured [1]. During this process, a certain type of sample holder or a specific way to hold the sample is employed. Due to the diverse characteristics of hydrated samples, such as water content, viscosity, strength, shape, or amount, various ways for sample holding are required. Consequently, more and more sample holders or approaches for holding samples are being developed. For example, Mo et al. developed a sample stage (holder) for the freeze-fracture of liquid, semi-liquid, and viscous samples, including chlorella [21]. Nonetheless, reliable protocols with easy and flexible sample-holding approaches that are compatible with various types of samples are still needed.
In this protocol, we propose an easy and reliable approach to hold and support the sample by employing a piece of filter paper as the substrate. The advantage of filter paper lies in its ability to support a thin film of sample, enabling a cold knife to make a cut and produce a clean freeze-fractured surface. This method is versatile and applicable to a broad range of samples, including cyanobacterial cells, algal cells, and any kind of sample that can be adsorbed onto a filter paper. Along with detailed, step-by-step operations for cryo-SEM tests, the authors believe that this protocol will be helpful for researchers to commence the design and operation of their cryo-SEM experiments.
Materials and reagents
Chlorella sorokiniana UTEX 1602 (Institute of Hydrobiology, Chinese Academy of Science, China, FACHB-275). In the following text, Chlorella sorokiniana UTEX 1602 is referred to as Chlorella sorokiniana
Filter paper (Hangzhou Special Paper Industry Co., Ltd, China, NEWSTAR) with a medium filtration speed. The thickness of the filter paper is 0.16 mm, and the pore size is approximately 7 μm. The diameter of a single piece of filter paper is 7 cm
Disposable transfer pipette (1 mL) (Jiangsu KangJian medical apparatus Co., Ltd, China, KANG JIAN, catalog number: KJ619)
Disposable sterile centrifuge tube (50 mL) (Hunan BKMAM Holding Co., Ltd, China, BKMAMLAB, catalog number: 20231206)
Liquid nitrogen (Dalian KeNa Technology Co., Ltd, China)
Equipment
Vacuum cryo transfer system (Leica, model: EM VCT500)
Field emission scanning electron microscopy (JEOL, model: JSM-7610 Plus)
Software and datasets
PC-SEM, v. 4.0.0.8 (Copyright 2006–2018 JEOL Ltd.)
Microsoft Office 2021 (Microsoft, https://www.office.com)
Jiangying Pro (v. 6.0.1, https://www.capcut.cn)
Procedure
Note: Leica EM VCT500 is a system consisting of several independent equipment, including the cryo-module equipped on SEM, which is referred to as VCT500; the Leica EM VCM for cryo-fixation of samples; the Leica EM ACE600 for freeze-fracture, freeze-etching, and sputter coating; and the shuttle to deliver the sample from different equipment in a vacuum and cryogenic state.
Preparation of the equipment
Mount the cryo-stage onto the scanning electron microscope (SEM) (see Figure 1, Video 1).
Loosen the spring snap of the airlock (Figure 1a).
Press the “vent” button on the front panel of the SEM to vent the chamber (Figure 1b).
While venting, unscrew the bolts on the chamber door (Figure 1c).
Pull open the chamber door carefully (Figure 1d).
Slide in the cryo-stage into the dovetail of the SEM stage (Figure 1e).
Set the type and height of the cryo-stage (Figure 1f).
Set the coordinates of the cryo-stage to the right position and angle (Figure 1h).
Mount the stopping piece next to the cryo-stage (red arrow in Figure 1j).
Mount the cold trap on top of the cryo-stage (Figure 1k).
Close the chamber door and evacuate the chamber (Figure 1l).
Figure 1. Step-by-step procedures of the mounting of cryo-stage. a. Loosening the spring snap of the airlock of the SEM chamber. b. Venting of the SEM chamber by pressing the “vent” button on the front panel. c. Unscrewing the bolts on the chamber door. d. Pulling open the chamber door. e. Sliding in the cryo-stage into the dovetail of the SEM stage. f. Setting the type and height of the cryo-stage. g. Mounted cryo-stage on the stage of SEM. h. Setting the coordinates of the cryo-stage. i. Cryo-stage in ready position for cryo-SEM experiment. j. Mounting the stopping piece next to the cryo-stage. k. Mounting the cold trap on top of the cryo-stage. l. Closing the chamber door.
Video 1. Mounting the cryo-stage into the dovetail of the SEM stage
Cool the cryo-stage of the vacuum cryo transfer system (VCT500), ACE600, and the shuttle (see Figure 2, Figure 3).
Fill liquid nitrogen into the dewar of the VCT500, ACE600, and the shuttle, as shown in Figure 2a-2, 2b-2, and 2c-2. Approximately 10 L of liquid nitrogen is required. To help understand the whole procedure, the VCT500, ACE600, and shuttle are shown in Figure 2a-1, b-1, and c-1, with red arrows indicating the main components.
Figure 2. Overview of the VCT500, ACE600, and the shuttle, along with the cooling processes. a-1. Control panel and cryo-dock of VCT500. a-2. Filling liquid nitrogen into the dewar of VCT500. b-1. ACE600 with red arrows indicating the dock, chamber, and handle of the cold knife. b-2. Filling liquid nitrogen into the dewar of ACE600. c-1. The shuttle attached to the VCM. c-2. Filling liquid nitrogen into the dewar of the shuttle. d. Mask and cryo-gloves for safety when handling liquid nitrogen.
Setting temperatures of the cryo-stage of VCT500 and ACE600.
Set the temperature of the cryo-stage of SEM on the control panel of VCT500 and activate the cooling, as shown in Figure 3a. Set the temperature of the cryo-stage of ACE600 and activate the cooling, as indicated in Figure 3b.
Figure 3. Setting the temperatures and the activation of cooling on the control panel of VCT500 (a) and ACE600 (b)
Load liquid nitrogen into VCM (see Figure 4b). Fill the reservoir of VCM with liquid nitrogen until 2/3 full.
Set the level of liquid nitrogen on the VCM (see Figure 4c). Select low level on the right of the control panel and press cooling to let liquid nitrogen flow from the reservoir to the liquid nitrogen bath.
Figure 4. Overview of the VCM and preparations for cryo-fixation. a. Picture of VCM with the shuttle attached. b. Filling liquid nitrogen into the reservoir of VCM. c. Control panel of VCM. d. Liquid nitrogen bath ready for cryo-fixation.
Preparation of the cryo-stub and application of Chlorella sorokiniana onto filter paper substrate
Fix the custom-designed sample holder onto the cryo-stub (Figure 5).
Put the pin of the holder into the hole in the middle of the cryo-stub (Figure 5b).
Tighten the screw in the cryo-stub to hold the pin in place (Figure 5c).
Adjust the holder in the right position on the cryo-stub so that in the consequent freeze-fracture procedure, the filter paper substrate fixed on the holder will be directly facing the cold knife (Figure 5d).
Figure 5. Setup of the custom-designed sample holder on the cryo-stub
Mount the filter paper substrate onto the sample holder.
Cut the filter paper into short strips with 3 mm × 5 mm.
Put the filter paper strips into the slots of the sample holder and tighten the screw to fix the filter paper (see Video 2).
Video 2. Fixation of the filter paper strips onto the sample holder
Apply a Chlorella sorokiniana droplet onto the filter paper (see Video 3). Use a 1 mL disposable transfer pipette to draw a small drop of water suspension of Chlorella sorokiniana. Put the tip against the top of the filter paper that is pre-fixed onto the sample holder. Apply the sample slowly onto the filter paper. Wash algal cells in distilled water by centrifugation at a relative centrifugal force of 3,754× g for 5 min.
Video 3. Appling a droplet of Chlorella sorokiniana onto the filter paper substrate
Cryo-fixation of the sample in liquid nitrogen bath (see Video 4)
Adjust the position of the cryo-pit inside the liquid nitrogen bath so that the cryo-stub could be slid in.
Use the handling rod to slide the cryo-stub quickly and steadily into the cryo-pit. The cryo-fixation time is approximately 3–5 min.
Video 4. Cryo-fixation of the sample in the liquid nitrogen bath
Transfer the cryo-stub from the VCM to ACE600
Hang the shuttle on the dock of VCM and press the attach button to attach the shuttle to VCM.
Retrieve the cryo-stub from the VCM to the shuttle (see Video 5).
Adjust the cryo-stub into the right position inside the cryo-pit.
Push the knob at the end of the transfer rod of the shuttle to let the holding-head approach the stub.
Insert the holding head inside the hole of the stub and turn the nob to 180° so the head can hold the stub tightly.
Pull the nob along the transfer rod to retrieve the cryo-stub back to the shuttle.
Press the detach button to pump the shuttle and close the valves, so the stub is in a vacuum and cryogenic state.
Video 5. Retrieving the cryo-stub from the VCM to the shuttle
Remove the shuttle from the VCM and attach it to the ACE600 (see Video 6).
Remove the shuttle from the dock of the VCM.
Hang the shuttle on the dock of the ACE600.
Press the attach button to attach the shuttle to the ACE600.
Video 6. Removal of the shuttle from the VCM and its attachment to the ACE600
Freeze-fracture of the sample
Transfer the cryo-stub from the shuttle to the cryo-stage of the ACE600 (see Video 7).
Push the knob at the end of the transfer rod of the shuttle to slide the cryo-stub onto the stage of ACE600.
Turn the nob to 180° to release the cryo-stub from the holding head.
Pull the nob back to the end of the transfer rod to retrieve the holding head.
Video 7. Transfer of the cryo-stub onto the cryo-stage of the ACE600
Freeze-fracture the sample (see Figure 6, Video 8)
Set the height of the cold knife. Select the freeze-fracture module on the control panel of the ACE600, as indicated in Figure 6a. Set the height of the cold knife (Figure 6b).
Figure 6. Setting the height of the cold knife. a. Selecting the freeze-fracture module from the main menu. b. Setting the height of the cold knife.
Video 8. Freeze-fracture of the samples with cold knife
Sweep the cold knife across the vertically standing samples and make a freeze-fracture (see Video 8). The cold knife is manipulated by a handle on the right side of the ACE600, as indicated by the red arrow in Figure 2b-1. The cold knife moves toward the samples and sweeps across the sample holder by pushing the handle forward. If one of the samples is not completely fractured, make one more sweep of the cold knife to force the upper part of the paper substrate to leave.
If the position of the cold knife is set too high above the sample holder, the filter paper will be bent instead of completely fractured. This is because the sample suspension film is thicker at the bottom and thinner at the top, where the filter paper might be too soft to be freeze-fractured. If the paper substrate is bent, as shown in Figure 7, the cross-sectional surface of the sample can still be revealed. Therefore, both the surface and cross-section of the sample are exposed at the same time.
Figure 7. Bent filter paper substrate showing both the cross-section and the surface
Clear the fracture debris on the sample (see Video 9): Pull the cryo-stub from the cryo-stage of ACE600 but keep it inside the chamber, and turn the stub to let the debris fall.
Slide the cryo-stub back to the cryo-stage of ACE600.
Video 9. Clearing of the debris after freeze-fracture
Freeze-etching and sputter coating
Set the freeze-etching parameters. Select the freeze-etching module from the main menu on the touchscreen of ACE600, as shown in Figure 8a. Set the temperature of etching, holding time, and ramp, as shown in Figure 8b.
Figure 8. Setting up the freeze-etching parameters. a. Selection of the freeze-etching module from the main menu. b. Setting up the temperature of etching, the holding time, and the ramp. c. Interface of freeze-etching.
Bring the cold knife right on top of the fractured sample (see Figure 9) and start the etching process, as indicated in Figure 8c.
Figure 9. Cold knife right on top of the sample during freeze-etching
Remove the cold knife from the top of the sample holder when the freeze-etching process ends.
Sputter coating: Select the sputter coating module from the main menu. Set the parameters of sputter coating and press the start button to initiate the sputter coating process, as shown in Figure 10.
Figure 10. Sputter coating. a. Setting up the parameters for sputter coating. b. Interface of sputter coating.
SEM investigation
Retrieve the cryo-stub from ACE600 back to the shuttle and detach the shuttle (see Video 10).
Retrieve the cryo-stub from ACE600 back to the shuttle by manipulating the transfer rod, as described in steps D2b–d.
Detach the shuttle from the ACE600.
Video 10. Retrieving the cryo-stub from ACE600 back to the shuttle
Attach the shuttle to the VCT500 and transfer the cryo-stub onto the cryo-stage of SEM (see Video 11).
Hang the shuttle on the dock of VCT500 and attach the shuttle.
Before starting to transfer the cryo-stub onto the cryo-stage of SEM, activate the camera on the PC-SEM software and set the cryo-stage in the right position.
When the attaching process is complete, push the knob carefully to slide the cryo-stub onto the cryo-stage of SEM.
Video 11. Transferring the cryo-stub from the ACE600 to the cryo-stage of SEM
Operation of SEM (see Video 12).
When the cryo-stub is transferred, adjust the height of it to a working distance of approximately 10 mm.
Turn on the electron beam.
By focusing and adjusting the contrast and brightness, the scattered Chlorella sorokiniana cells in ice can be easily found. The freeze-fractured surface is flat and wide enough to get adequate spots of interest.
Video 12. SEM operation
Ending procedures
Remove the cryo-stub from the SEM (see Video 13).
Set the cryo-stub into the right position for transfer.
Carefully transfer the cryo-stub from the cryo-stage of the SEM back to the shuttle.
Detach the shuttle from SEM and attach it to the VCM.
Transfer the cryo-stub from the shuttle to the cryo-fixation pit of VCM.
Remove the cryo-stub from the pit and put it on the hot plate.
Video 13. Retrieving the cryo-stub from the SEM back to the shuttle
Bake out all the equipment, including VCT500, ACE600, VCM, and the shuttle (see Figure 11).
Press the bake out button on the main menu.
Set the bake out time.
Execute the bake out procedure.
Remove all the lids on the dewar.
Figure 11. Baking out all the equipment. a. VCT500. b. ACE600. c. VCM and the shuttle.
Data analysis
The data obtained by this protocol is mainly SEM images, and the data bar is incorporated using Microsoft PowerPoint.
Validation of protocol
The cryo-SEM experiment following this protocol was performed five times, and each replicate showed similar results concerning the freeze-fracture process and the SEM results. SEM images are shown in Figure 12. The images show a clean fractured surface with algae cells embedded in ice (Figure 12a); some cells are cleaved, revealing the internal structure, as shown in Figure 12a-2 and 12a-3. The sample that is etched after freeze-fracture showed scattered algae cells standing on top of the ice, revealing a clearer profile of the cell (Figure 12b). The parameters of freeze-etching are included in general notes.
Figure 12. SEM images of freeze-fractured samples showing the surface and internal structure. a. Fractured sample of Chlorella sorokiniana without etching. b. After etching.
General notes and troubleshooting
Before venting the SEM chamber, make sure to loosen the spring snap of the airlock; otherwise, the nitrogen flow introduced into the chamber may lead to overpressure and damage the film window of the EDS detector.
When setting the type and height of the cryo-stage, it is recommended to choose the 32 mm holder and set the height as 20 mm to avoid collision with the pole piece.
When mounting the cryo-stage onto the SEM, it is recommended to mount the cryo-stage a day before the cryo-SEM experiments, since the evacuation time could take more than 3 h.
Be cautious when handling liquid nitrogen and make sure to wear a mask and cryo-gloves, as shown in Figure 2d.
As for the temperature of the cryo-stage of SEM, it is recommended to set it to -140 °C or -150 °C. A higher temperature may lead to sample sublimation and contamination of the SEM chamber. As for the temperature of the cryo-stage of ACE600, if a freeze-etching process is performed right after the freeze-fracture, set it to the starting temperature of the etching process. If no freeze-etching is performed, set the temperature to that of the cryo-stage of SEM.
The custom-designed sample holder is made for holding thin film samples vertically so that they can be easily freeze-fractured. The holder is made of copper and both sides are able to hold samples.
The holding head of the transfer shuttle is the component located at the head of the transfer rod. It plays the role of holding the stub during the transfer process. It is made of special resin and is half-transparent.
Before loading the cryo-stub onto the cryo-stage of ACE600, it is important to check the stage temperature prior.
The clearing of the freeze-fractured samples is necessary because it could get rid of debris in the subsequent procedures and avoid the contamination of the SEM chamber.
The freeze-etching procedure plays an important role in getting fine details of the inner structure. Under-etching or over-etching may either cause false surface morphology or damage the Chlorella sorokiniana cells. In this protocol, the freeze-fractured sample is investigated with and without freeze-etching. The freeze-etching parameters are as follows: The temperature of the cryo-stage is -100 °C, the etching temperature is -85 °C, the holding time is 40 min, and the ramp is 5 °C/min. Note that the difference in temperature between the starting temperature and the etching is ideally 15–20 °C.
The cold knife plays as a cold trap during the etching process, so it is recommended to place it as close to the surface of the sample as possible.
Before sputter coating, make sure to remove the cold knife from the top of the samples. The metal used for sputter coating in this experiment is Pt. The ACE600 is equipped with a quartz monitor to record the thickness of Pt. Both filter paper and ice are materials with poor conductivity, so the sputter time is set to 120 s and the thickness of Pt is approximately 5 nm.
The 5-axis coordination of the cryo-stage for sample transfer was set by the supplier upon the installation of the VCT500.
The accelerating voltage of the electron beam in SEM investigation is recommended to be below 3 kV, since an electron beam with higher energy may cause severe surface damage and charging effect.
Frost is one of the primary obstacles to a successful cryo-SEM experiment. It can be a thick layer of nanocrystals covering the sample features, and it can get the valves and cryo-stub stuck during the transfer process. Therefore, operators should minimize the frosting on samples and components of the equipment.
When pulling the cryo-stub from the liquid nitrogen bath of VCM, it will contact with the air for a few seconds. Although VCM has a patent to avoid the formation of frost during this process, it is important that operators perform this transfer as quickly as possible
Be aware that when attached to the VCM, the shuttle is connected to ambient air. A long time of exposure to air will cause severe frosting on the cryo-stage and valves. Make sure to detach the shuttle from VCM whenever the transfer process is done. When transferring the sample from the VCM, only attach the shuttle when the sample is ready for transfer.
Keep in mind that the sample should be immersed in liquid nitrogen throughout the whole cryo-fixation process.
Tools like tweezers and handling rods should be placed on the hot plate to be heated up immediately after being used in liquid nitrogen. They are ready for use again only when the frost is completely gone.
Use liquid nitrogen free of ice crystals, especially the liquid nitrogen used for the LN2 bath for cryo-fixation in the VCM.
The bake out is the process of heating the dewar and the inner components of VCM, ACE600, VCT500, and the shuttle to remove all the frost, water, and contaminants. After the bake-out process, the equipment is dry, clean, and ready for use.
It is recommended to set the bake-out time to maximum. Otherwise, the residual water could turn into ice and get the components stuck in the consequent cryo-SEM experiment.
A sputter-coated sample can technically be freeze-etched to remove excessive ice from the surface, but it is not recommended. A metal film a few nanometers thick could change the freeze-etching process of ice drastically, and the metal film would be obvious on the surface when the ice beneath it is gone.
Carbon tape is not a good option to adhere samples onto stubs and holders in cryo-experiments; use toothpaste or saturated sucrose solution instead.
Acknowledgments
This work was funded by the Foundation of Instrumental Maintenance and Research by Dalian University (No. SYSWX202301).
Competing interests
The authors declare no competing financial interests.
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Article Information
Publication history
Received: Jul 24, 2024
Accepted: Oct 14, 2024
Available online: Nov 4, 2024
Published: Dec 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Microbiology > Microbial cell biology > Cell imaging
Cell Biology > Cell imaging > Electron microscopy
Biophysics > Electron cryotomography
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5,144 | https://bio-protocol.org/en/bpdetail?id=5144&type=0 | # Bio-Protocol Content
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Peer-reviewed
Temporally and Spatially Controlled Age-Related Prostate Cancer Model in Mice
SL Sen Liu
KS Keyi Shen *
ZL Zixuan Li *
SR Seleste Rivero
QZ Qiuyang Zhang
(*contributed equally to this work)
Published: Vol 15, Iss 1, Jan 5, 2025
DOI: 10.21769/BioProtoc.5144 Views: 242
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Original Research Article:
The authors used this protocol in Frontiers in Molecular Biosciences Jun 2021
Abstract
The initiation and progression of prostate cancer (PCa) are associated with aging. In the history of age-related PCa research, mice have become a more popular animal model option than any other species due to their short lifespan and rapid reproduction. However, PCa in mice is usually induced at a relatively young age, while it spontaneously develops in humans at an older age. Thus, it is essential to develop a method by which the PCa initiation and progression timeline can be strictly controlled to mimic human physiological conditions. One milestone in this field was the identification of the prostate-specific transcription factor, Probasin (Pb), which allowed for the prostate-specific expression of genes knocked into the mice's genome. Another milestone is the establishment of the preclinical mouse model with Pten conditionally knocked out in the prostate tissue, which closely mimics the formation and growth of human PCa. Hereby, we present the prostate-specific temporally and spatially controlled Pten knockout PCa mouse model that can be induced using an adenovirus-based Cre-LoxP system. The Cre recombinase (Cre) is inserted into an adenovirus vector. Unlike Pb-Cre knock-in models (which are spatially but not temporally controlled), the expression of Cre is activated to knock out Pten from the mice's prostate epithelial cells once injected. The viral delivery procedures strictly control the location and time of Pten knockout. This novel approach provides a powerful age-related murine model for PCa, emphasizing the effect of aging on prostate carcinogenesis.
Key features
• In vivo delivery of Cre recombinase adenovirus (Ad-Cre-Luc) in Pten LoxP/LoxP (L/L) mice.
• Generation of Cre-expressing Ad-Cre-Luc-mediated ablation of Pten in anterior prostate epithelial cells of adult Pten L/L mice at different ages.
• The Ad-Cre-Luc-mediated ablation of Pten leads to hyperplasia that progresses through prostatic intraepithelial neoplasia (PIN) to adenocarcinoma.
• PIN refers to the non-cancerous growth of epithelial cells in the prostate tissue—not cancer but a precursor of prostate cancer [1].
Keywords: In vivo viral delivery Adeno-Cre-Luc virus Pten conditional knockout mice Prostate cancer animal model
Graphical overview
Surgery, luciferase-based bioluminescent imaging, and histological observation
Background
Among all risk factors, age is the most important one associated with prostate cancer (PCa) [2]. Canine models are considered valuable in preclinical studies of PCa because they are similar to humans in terms of size, anatomy, histology, and spontaneity of PCa [3]. However, the relatively long lifespan and low incidence rate may limit the use of these models, especially in age-related studies [3]. In contrast, mice's rapid reproduction and short lifespan make them ideal models in which the role of aging in PCa can be studied relatively quickly. However, PCa in mice models must be induced, which is less likely to resemble human conditions, especially the age of PCa initiation. Hence, there is an urgent need to develop a murine model where the timeline of PCa initiation and progression can be strictly controlled.
A milestone in this field is the creation of the prostate-specific Pten conditional knockout (cKO) mouse model, which closely mimics the formation and growth of human PCa and has become an established PCa preclinical model [4]. However, as Pten deletion in this model is an automatic process triggered in very young prostate tissue (less than eight weeks old), it is challenging to mimic the physiology and the microenvironment (old prostate tissue) in which human PCa develops. We created a novel mouse model where Pten KO can be spatially and temporally controlled (Pten adcre+) using a virus-assisted in vivo cKO approach. The prostate-specific Cre-LoxP gene switching was generated via intraductal delivery of adenovirus to the anterior prostate lobes [5]. Adenovirus is a DNA virus that does not integrate into the host genome. It infects dividing and nondividing cells, leading to a transient high-level protein expression [6]. Using the intraductal delivery method, we obtained epithelial-specific prostate infection with Cre-expressing adenoviruses, leading to the deletion of the floxed Pten gene in the prostate epithelium.
Materials and reagents
Biological materials
1. Ptenflox mice (The Jackson Laboratory, strain number: 006440)
2. Cre recombinase adenovirus (Ad-Cre-Luc) (VECTOR BIOLABS, catalog number: 1705); co-expression of Cre recombinase and luciferase (the reporter gene) is regulated by the cytomegalovirus (CMV) promoter
3. Luciferase adenovirus (Ad-CMV-Luc) (VECTOR BIOLABS, catalog number: 1000); this adenovirus vector delivers luciferase. Expression is also regulated by the CMV promoter. It is used as a Cre-negative control of the Ad-Cre-Luc viral vector
Reagents
1. Lidocaine 2% (VETONE, catalog number: V1 510212)
2. OstiLOX (VETONE, catalog number: V1 501080)
3. Sodium chloride (NaCl) (Fisher Chemical, catalog number: S271-10)
4. 10% povidone iodine (McKesson, catalog number: MFR#036)
5. Paraformaldehyde (Sigma-Aldrich, catalog number: P6148-1KG)
6. 100% ethanol, anhydrous (Fisher Chemical, catalog number: A405P-4)
7. Xylene (Thermo Scientific, catalog number: 9990501)
8. CAT hematoxylin (BIOCARE MEDICAL, catalog number: CATHE-GL)
9. Edgar degas eosin (BIOCARE MEDICAL, catalog number: THE-GL)
10. Tacha's bluing solution 10× (BIOCARE MEDICAL, catalog number: HTBLU-MX)
11. Saturated HCl (Fisher Scientific, catalog number: 60047420)
12. Soothe eye ointment (Bausch & Lomb, Preservative Free, 1.8 Oz)
13. Blocking serum from VECTASTAIN Elite ABC Kit (VECTOR LABORATORIES INC, catalog number: PK-6101); see Recipes
14. Biotinylated secondary antibody from VECTASTAIN Elite ABC Kit (VECTOR LABORATORIES INC, catalog number: PK-6101); see Recipes
15. 3,3'-diaminobenzidine (DAB) solution (VECTOR LABORATORIES INC, catalog number: sk-4100); see Recipes for final concentration
16. Phosphate-buffered saline (PBS) (GENESEESCI, catalog number: 25-508)
17. 30% hydrogen peroxide (H2O2) (Fisher Scientific, catalog number: H254-500)
18. D-luciferin (Cayman Chemical Company, catalog number: 14682)
Solutions
1. OstiLox for injection (see Recipes)
2. Lidocaine for injection (see Recipes)
3. 10% paraformaldehyde (see Recipes)
4. 15% ethanol (see Recipes)
5. 70% ethanol (see Recipes)
6. 80% ethanol (see Recipes)
7. 95% ethanol (see Recipes)
8. Bluing solution (see Recipes)
9. 0.5% HCl-ethanol solution (see Recipes)
10. Blocking serum from VECTASTAIN Elite ABC Kit (see Recipes)
11. Biotinylated secondary antibody from VECTASTAIN Elite ABC Kit (see Recipes)
12. Avidin-Biotin-peroxidase Complex (ABC) solution from VECTASTAIN Elite ABC Kit (see Recipes)
13. DAB solution (see Recipes)
Recipes
1. OstiLox for injection
Reagent Final concentration Quantity or Volume
OstiLox 1 mg/mL 200 μL
NaCl 9 mg/mL 800 μL
Total n/a 1 mL
2. Lidocaine for injection
Reagent Final concentration Quantity or Volume
Lidocaine 2% 4 mg/mL 200 μL
NaCl 9 mg/mL 800 μL
Total n/a 1 mL
3. 10% paraformaldehyde
Reagent Final concentration Quantity or Volume
Paraformaldehyde 100 g/L 100 g
PBS 1× 1 L
Total n/a 1 L
4. 15% ethanol
Reagent Final concentration Quantity or Volume
Ethanol anhydrous 15% (v/v) 150 mL
Deionized H2O n/a Adjust the total volume to 1 L
Total n/a 1 L
5. 70% ethanol
Reagent Final concentration Quantity or Volume
Ethanol anhydrous 70% (v/v) 700 mL
Deionized H2O n/a Adjust the total volume to 1 L
Total n/a 1 L
6. 80% ethanol
Reagent Final concentration Quantity or Volume
Ethanol anhydrous 80% (v/v) 800 mL
Deionized H2O n/a Adjust the total volume to 1 L
Total n/a 1 L
7. 95% ethanol
Reagent Final concentration Quantity or Volume
Ethanol anhydrous 95% (v/v) 950 mL
Deionized H2O n/a Adjust the total volume to 1 L
Total n/a 1 L
8. Bluing solution
Reagent Final concentration Quantity or Volume
Tacha's bluing solution 10× 1× 100 mL
Deionized H2O n/a Adjust the total volume to 1 L
Total n/a 1 L
9. 0.5% HCl-ethanol solution
Reagent Final concentration Quantity or Volume
Saturated HCl 0.5% (v/v) 1 mL
70% ethanol 1× 199 mL
Total n/a 200 mL
10. Blocking serum from VECTASTAIN Elite ABC Kit
Reagent Final concentration Quantity or Volume
Blocking serum 1.5% (v/v) 150 μL
PBS 1× 10 mL
Total n/a 10.15 mL
11. Biotinylated secondary antibody from VECTASTAIN Elite ABC Kit
Reagent Final concentration Quantity or Volume
Biotinylated antibody 1:200 diluted 50 μL
Blocking serum 1.5% (v/v) 150 μL
PBS 1× 10 mL
Total n/a 10.2 mL
12. ABC solution from VECTASTAIN Elite ABC Kit
Reagent Final concentration Quantity or Volume
Avidin DH 1:50 diluted 100 μL
Biotinylated horseradish peroxidase H 1:50 diluted 100 μL
PBS 1× 5 mL
Total n/a 5.2 mL
13. DAB solution
Reagent Final concentration Quantity or Volume
PBS 0.0152× 15.2 μL
DAB 0.5 g/L 17.6 μL
H2O2 0.15 g/L 16 μL
Deionized H2O n/a Adjust the total volume to 1 mL
Total (optional) n/a 1 mL
Laboratory supplies
1. Microscope slides (Fisher Scientific, catalog number: 12-550-15)
2. Cover glass (Brain Research Laboratories, catalog number: 2222-1.5D)
3. Tissue cassettes with recessed cover (Simport Scientific, catalog number: M509-2)
4. 4-Ply non-woven, non-sterile dental gauze sponges (PlastCare USA, catalog number: PG‐3304‐3)
5. GLAD PRESS'N SEAL food wrap (Phoenix Research Industries INC, catalog number: CLO70441)
6. 5-0 nylon sutures (Securos Surgical, catalog number: CG634)
7. SimPort tissue cassette (SimPort Scientific INC, catalog number: M-509-2)
8. ImmEdge pen (VECTOR LABORATORIES INC, catalog number: H-4000)
9. Coverslip 22 × 22 (Fisher Scientific, catalog number: C10228)
Equipment
1. Bead sterilizer: The germ terminator (CellPoint scientific, model: GERMINATORTM 500)
2. Biological safety cabinet (The Bader Company, catalog number: SG404)
3. Surgipath Paraplast surgical microscope (Leica, catalog number: 39601006)
4. Gas anesthesia system (Caliper LifeScience, model: XGI-8)
5. IVIS-Lumina (Caliper LifeScience, model: XRMS)
6. Microtome (New Life Scientific INC, model: HM355S)
7. Isotemp incubator (Fisher Scientific, model: 637D)
8. Nikon microscope (PRECISION INSTRUMENTS, LLC, model: H550S)
9. 50 μL, model 705 RN syringe (Hamilton, catalog number: CAL7637-01)
10. 33 gauge, small hub RN needle, custom length (0.5 to 12 in.), point style 2, 3 or 4, 6/PK (Hamilton, catalog number: 7803-05)
11. Decloaking ChamberTM NxGen (BIOCARE MEDICAL, model: DC2012)
12. Sterile surgical tools (forceps, scalpel, small scissors, and hemostats)
13. Micropipette (Hamilton, catalog number: 7637-01)
Software and datasets
1. PureCLIP (Sabrina Krakau et al., https://github.com/skrakau/PureCLIP) [7]
2. NIS-Elements BR Analysis (Nikon, 4.60.00, https://www.microscope.healthcare.nikon.com/products/software/nis-elements/nis-elements-basic-research)
Procedure
A. Surgery (Day 1)
1. Retrieve frozen viral vectors (Ad-Cre-Luc and Ad-CMV-Luc, see Biological materials) from the -80 °C freezer. Keep the virus on ice.
Note: Cre recombinase catalyzes the site-specific DNA recombination between LoxP sites. The adenovirus titer is 1 × 1010 PFU/mL.
2. Locate and identify the mice before the surgery using the cage card information and ear puncture code.
3. Record the weight, age, and sex.
4. Inject a mouse with an analgesic drug (OstiLOX, subcutaneous injection, SC). The dosage is calculated based on the mouse's weight (0.1 mg/kg).
5. Remove the hair using an electric shaver. Avoid potential thermal or physical lesions caused by the fast-acting hot blade. Note that the blades are hot because they are rubbing against each other (like scissors).
6. Clean the area shaved in the previous step with tape (optional).
7. Turn on the heat of the operating table before anesthetizing the mouse so that it is warm enough at the beginning of the surgery to prevent potential hypothermia.
8. Turn on the oxygen and adjust the anesthesia machine to a flow rate of 0.4 L/min (Figure 1A).
Figure 1. Surgical and imaging procedures. (A) Anesthetize the mouse. (B) Cover the mouse under a semi-transparent food wrap with the surgical field exposed (outlined with dashed lines). (C) Create an abdominal incision using a scalpel. (D) Expose the genitourinary (GU) bloc. The cartoon and the magnified photo of the mouse's GU bloc demonstrate the anatomy of seminal vesicles, anterior prostate tissue, and the urinary bladder. L, left. R, right. AP, anterior prostate. SV, seminal vesicles. (E) Inject Ad-Cre-Luc viral particles into APs on both sides (this panel only shows injection into the right AP). Ad-Cre-Luc, adenovirus viral particle driving the co-expression of Cre recombinase and luciferase. (F) Close the incision. (G) Let the mouse recover from anesthesia. (H) Anesthetize the mouse. (I) Peritoneal injection of D-luciferin. (J) Detect the infection of Ad-Cre-Luc using luciferase-based imaging. (K) Let the mouse recover from anesthesia.
9. Anesthetize the mouse with 2%–4% isoflurane in oxygen.
10. Once the mouse is anesthetized in the recovery cage, move it onto the operating table, leaving it supine. Then, place the nose cone over the snout and turn the isoflurane flow rate to 2%.
11. Apply the lubricant eye ointment to the mouse's eyes bilaterally to prevent corneal dehydration.
12. Determine the depth of anesthesia by the pedal withdrawal (toe-pinch) reflex test. Once the reflex is tested negative, turn the isoflurane flow rate down to 1%. Check the respiratory rate and responsiveness to toe pinch every 5 min.
13. Use alternating swabs of 10% povidone iodine and 70% ethanol to wipe the abdominal area three times.
14. Cover the mouse with a clear food wrap (Figure 1B).
15. Cut a hole in the food wrap to expose the desired operating field (in the abdominal area, Figure 1B).
16. Before making the incision, inject the mouse (intradermal injection, ID) with local anesthetics (lidocaine 2%).
17. Confirm there is no toe pinch response.
18. Use the scalpel to cut a midline incision down the sagittal plane of the mouse (Figure 1C). Use the small scissors to enlarge the incision to the desired length (approximately 1.5–2 cm). Use the hemostatic forceps to grab the midpoint of the skin on each side of the incision to expose the operating field.
19. Upon opening the abdomen, look for a round, semi-transparent structure in the caudal abdomen or the pelvic region. That is the mouse's urinary bladder. Use it as a landmark for the upcoming procedures (Figure 1D).
20. Use a tweezer to pick up the bladder. Look for seminal vesicles in the immediate vicinity of the urinary bladder. The seminal vesicles are symmetrical, pale-yellow, curved or spiral-shaped, elongated tubular glands with a sac-like appearance (Figure 1D).
21. Lift the seminal vesicles gently using tweezers. Be very careful because the highly vascularized anterior prostate (AP) tissues are attached to seminal vesicles.
22. Follow the arteries running along the seminal vesicles. As they approach the base of the bladder, they are attached to the APs, the glandular structures with a pale and smooth surface (Figure 1D).
23. Place the anesthetized mouse on a heating pad and move it into the biosafety cabinet (BSC).
24. Under a surgical microscope, look for the primary ducts of the APs. These are the sites where the virus will be delivered.
25. Intraductally deliver 5–10 μL (4–8 × 106 PFU/g of body weight) of viral solution (1 × 1010 PFU/mL) into the AP using a micropipette equipped with a 27 G needle (Figure 1E).
26. Place the organs back to their original anatomical positions.
27. Suture the muscular abdominal wall and the skin individually using 5-0 nylon sutures (Figure 1F).
28. After the surgery and injection described above, observe the mouse until it is awake and able to walk independently. Keep the mouse on a warm surface and monitor its respiratory rate until complete recovery from anesthesia (Figure 1G).
29. If available, repeat the surgical procedure described above on different mice (with the same genotype, Pten L/L).
B. Luciferase-based bioluminescent imaging (Days 5–7)
During the window of days 5–7, the mice are subjected to luciferase-based bioluminescent imaging using the IVIS-Lumina XRMS in vivo imaging system.
1. Apply eye ointment to the mouse's eyes to prevent corneal dehydration.
2. Anesthetize mice with 2%–4% isoflurane in oxygen (Figure 1H).
3. Inject 10 μL of D-luciferin (15 mg/mL sodium salt, Figure 1I) intraperitoneally per gram of body weight. For instance, give a 20 g mouse approximately 200 μL for a standard 150 mg/kg injection.
4. Start image acquisition immediately with a series of images within 30 min (Figure 1J). The peak light emission represents the expression of injected Ad-Cre-Luc infection (Figure 2).
5. After image acquisition, observe the mouse until it is awake and able to walk independently. Keep the mouse on a warm surface and monitor its respiratory rate until complete recovery from anesthesia (Figure 1K).
Figure 2. Luciferase-based bioluminescent imaging. Representative image of a mouse with bilateral anterior prostate (AP) infection of Adenovirus that drives the expression of the reporter gene, luciferase. The image was acquired five days after the surgery. The color-coded scale bars on the right side indicate signal intensities of the luciferase-catalyzed reaction in the image on the left.
C. Tissue harvesting
After the surgery, tissue harvesting takes place at 4, 8, and 16 weeks.
1. Euthanize the mouse by carbon dioxide asphyxiation.
2. Pin the mouse on a dissection board in a supine position by poking a pin through each of the four paws.
3. Spray the abdominal surface with 70% ethanol.
4. Hold the skin of the lower abdomen with tweezers and use scissors to cut through the skin and the abdominal wall, approximately 1 cm anterior to the opening of the penis. Be careful not to cut any organs.
5. Enlarge the incision created in the previous step, exposing the peritoneal cavity.
6. Identify the bladder and urogenital tract and expose them by gently moving fat and other organs to the side.
7. Using tweezers, grip the vas deferens at the base near the urethra and tear it away.
8. Repeat the previous step on the contralateral vas deferens.
9. Carefully lift the bladder with tweezers. Meanwhile, use scissors to cut the urethra below the bladder and ventral prostate.
Note: As the bladder is pulled out, the entire genitourinary (GU) bloc, except the testis, will come with it (Figure 3A).
Figure 3. Tissue harvesting. (A) The GU bloc is separated from the rest of the mouse. It is placed in the same orientation as it is in a mouse in the supine position. The ruler uses a metric unit (centimeter). APs are outlined with dashed circles. GU bloc, genitourinary bloc. SV, seminal vesicle. AP, anterior prostate. (B) The right AP is being separated from the right seminal vesicle. L, left. R, right.
10. Place the GU bloc in a 60 mm cell culture dish containing 5–10 mL of PBS.
11. Perform the remaining part of the protocol under a dissection microscope. Use two pairs of fine tweezers, one in each hand. Hold the tweezers like a pencil and rest the forearms on the benchtop to stabilize them.
12. Use fine tweezers to position the GU bloc so that the bladder is on top, the urethra is pointing down, and the seminal vesicles are placed on either side of the bladder (Figure 3A).
13. Grip the urethra with tweezers and carefully pull away fat with the other forceps without tearing away any prostate tissue.
Note: Fat will appear "shiny" relative to the surrounding tissue.
14. Once the fat is removed, use the tweezers to tear the connective tissue between the two ventral lobes of the prostate and separate them.
15. To remove one of the anterior lobes of the prostate (AP), use tweezers to gently tear the lobe away from the seminal vesicle, ensuring not to puncture the seminal vesicle (Figure 3B). When the lobe is apart from the seminal vesicle but still attached to the base of the urinary bladder, cut it away with scissors.
16. Repeat step C15 with the contralateral AP.
17. Place the APs in Eppendorf tubes. Label the tubes and place them in liquid nitrogen to freeze the tissue.
18. Fix the remainder of the GU bloc with 10% paraformaldehyde.
D. Tissue processing
1. Treat the tissue (GU bloc) with ethanol in an increasing concentration step by step. Use a tilting stand to shake the tissue container so the ethanol can move around the tissue. Ethanol concentrations are 70%, 80%, 95%, and 100%. Each step takes 1 h. Repeat the step with fresh ethanol when the concentration is ≥95% (Table 1). The purpose of the serial ethanol treatments is to dehydrate the tissue gently. Trim the tissue after the 80% ethanol treatment by cutting away part of the seminal vesicles so that the ethanol can infiltrate the tissue more effectively.
2. Transfer the dehydrated tissue to glass vials. Treat the tissue with xylene.
Table 1. Procedures for tissue processing
Step Temperature Duration Comment
70% ethanol Room temperature 1 h Tissues stay in the same container in which they were fixed with PFA.
80% ethanol Room temperature 1 h Trim the tissue after this step.
95% ethanol I Room temperature 1 h
95% ethanol II Room temperature 1 h
100% ethanol I Room temperature 1 h
100% ethanol II Room temperature 1 h
Xylene I Room temperature ≤ 5 min Move the tissues to glass vials.
Xylene II Room temperature Varies Depending on when the tissue turns semi-transparent.
E. Tissue embedding
1. Place the tissue from the previous day (fixed in wax) in an incubator. Set the temperature to 65 °C and let the wax melt.
2. Meanwhile, let fresh wax melt at 65 °C.
3. Transfer the tissue from melted wax to an empty mold.
4. Adjust the tissue's position so that the cross-section of the mouse's urethra is facing down.
5. Fill the mold with melted fresh wax.
6. Place a SimPort® tissue cassette on top of the mold. Add more wax so the tissue cassette can hold the entire block once the wax solidifies.
7. Place the entire block (a mold containing tissue and wax with a tissue cassette attached to it) on a cold table surface.
8. Once the surface of the wax solidifies, place the entire block into cold tap water (no need to use distilled or deionized water).
9. Store the solidified blocks in a -20 °C freezer.
F. Tissue sectioning
1. Trim the blocks with a blade so that the volume of wax is minimized but still able to hold the tissue.
2. Load the microtome with blocks from the previous step.
3. Adjust the position of the block. Make sure the mouse's urethra is perpendicular to the blade.
4. Program the microtome to make it cut sections with a thickness of 4 μm.
5. Let the microtome cut four consecutive sections automatically.
6. Carefully pick up the consecutive sections with tweezers. Place the consecutive sections on the surface of 15% ethanol to reduce the wrinkles.
7. Place a fresh slide in the water bath underneath the floating sections. Approach the sections quickly but carefully so that the wrinkles can be fully reduced, but the sections can be picked up before the wax melts. Let one side of the consecutive sections cling to the slide, and then pick up the slide carefully. Make sure the tissue in all four sections stays on the slide.
8. Place the slide on a warm surface and let it dry.
G. Immunohistochemistry (IHC) and hematoxylin & eosin (H&E) staining
Steps G1–7 are summarized in Table 2.
1. Place the slides in an incubator. Set the temperature to 65 °C. Incubate for 30 min.
2. Transfer to a xylene bath and incubate for 5 min.
3. Transfer to a fresh xylene bath and incubate for another 5 min.
4. Shake off excess liquid and treat the tissue with two rounds of 100% ethanol (3 min each).
5. Shake off excess liquid and treat the tissue with two rounds of 95% ethanol (3 min each).
6. Shake off excess liquid and place slides in fresh 80% ethanol for 3 min.
7. Shake off excess liquid and place slides in fresh 70% ethanol for 3 min.
Table 2. Wax removal and rehydration
Step Temperature Duration Comment
Wax removal 65 °C 30 min Wax is liquified at 65 °C.
Xylene I Room temperature 5 min Liquified wax is dissolved in the xylene.
Xylene II Room temperature 5 min A second round of xylene treatment cleans up residual wax.
100% ethanol I Room temperature 3 min Absolute ethanol dissolves xylene.
100% ethanol II Room temperature 3 min A second round of ethanol cleans up residual xylene.
95% ethanol I Room temperature 3 min Gentle rehydration with ethanol in a series of decreasing concentrations.
95% ethanol II Room temperature 3 min
80% ethanol Room temperature 3 min
70% ethanol Room temperature 3 min
8. Rinse the slides in gently running tap water for 30 s (avoid a direct jet, which may wash off the section).
9. Place in a PBS wash bath for further rehydration (30 min at room temperature).
10. Turn on the power for the decloaking chamber. Set the temperature to 95 °C and set a timer for 5 min.
11. Let the decloaking chamber pre-heat. Meanwhile, load the slide holder with slides and place the slide holder in a container (provided by the manufacturer of the decloaking chamber) filled with 0.1 mM EDTA solution. Make sure the tissue is completely submerged in the EDTA solution.
12. Once the decloaking chamber reaches 95 °C, place the EDTA container (from the previous step) in it. Let the reaction start.
13. At the end of the 5-min countdown, remove the entire EDTA container and let it cool to room temperature.
14. Discard the EDTA and rinse the slides with tap water (do NOT use distilled or deionized water).
15. Place the slides on a horizontal surface. Do not allow the slides to touch each other. Between each step, keep the tissue hydrated with PBS.
16. Draw circles around the tissue with an ImmEdgeTM Pen. The "ink" is a hydrophobic substance so that the circles around the tissue can contain aqueous solutions within the area.
17. Add drops of 3% hydrogen peroxide to cover all the consecutive sections.
18. Incubate at room temperature for at least 10 min.
19. Wash the tissue three times with PBS (5 min each).
20. Shake off the fluid with a brisk motion and carefully wipe each slide around the sections.
21. Apply blocking serum.
22. Dilute the primary antibodies in the same blocking serum.
23. Apply the primary antibody solution to the tissue sections, ensuring sufficient antibody solution covers each section. In addition, watch for the integrity of the circles drawn in step G16 so that different antibodies do not cross-contaminate each other. Re-draw the circles if necessary.
24. Incubate overnight at 4 °C.
25. At this moment, one of the four consecutive sections must be covered in PBS and reserved for Hematoxylin & Eosin (H&E) staining in the future.
26. Remove the primary antibody from the previous step.
27. Wash the tissue three times with PBS (5 min each) and keep the tissue covered in PBS.
28. Prepare the biotinylated secondary antibody using the VECTASTAIN® Elite ABC Kit (see Recipes).
29. Apply the secondary antibody. Incubate for 1 or 2 h at room temperature.
30. During the last 30 min of the secondary antibody reaction, start preparing the avidin/oxidizable peroxidase reagent using the VECTASTAIN® Elite ABC Kit (see Recipes).
31. Meanwhile, prepare the 3,3'-Diaminobenzidine (DAB) solution as described in the recipe. The addition of each reagent must follow the order listed in the recipe. Otherwise, a precipitate could form.
32. Remove the biotinylated secondary antibody and wash three times with PBS (5 min each).
33. Apply the ABC solution and incubate at room temperature for 30 min.
34. Remove the ABC solution and wash three times with PBS (5 min each).
35. Apply the DAB solution to develop color.
36. Remove the DAB solution and wash three times with PBS (5 min each).
37. Place the slides on a slide holder.
38. Place the slide holder from the previous step in a hematoxylin bath for 3 min.
39. Rinse the slides with tap water.
40. Dip the slides a few times in an ethanol-based 0.5% (v/v) HCl solution (see Recipes) to remove the excess hematoxylin.
41. View the slides under a microscope. Based on the darkness of the nuclei, decide whether hematoxylin or HCl/ethanol treatment needs to be repeated.
42. Place the slides in a bath of bluing solution for 3 min.
43. Keep the IHC sections covered in PBS. Apply eosin to the section reserved for H&E staining.
44. Dehydrate the tissue with ethanol in order of increasing concentrations. This procedure reverses the rehydration process. Instead of using an ethanol bath, apply drops of ethanol to each section separately to prevent cross-contamination (Table 3).
Table 3. Dehydration and mounting of coverslip
Step Temperature Duration Comment
70% ethanol Room temperature 3 min Tissues stay in the same container in which they were fixed with PFA.
80% ethanol Room temperature 3 min Trim the tissue after this step.
95% ethanol I Room temperature 3 min
95% ethanol II Room temperature 3 min
100% ethanol I Room temperature 3 min
100% ethanol II Room temperature 3 min
Xylene I Room temperature 3 min
Xylene II Room temperature 20 min
Resin and coverslip Room temperature Overnight Add a few drops of xylene so that it is easier to spread the resin across the slide.
H. Image acquisition
1. Turn on the Nikon Digital Sight DS Fi1 Camera using the power button.
2. Open the NIS-Elements Basic Research Software to connect the microscope and camera.
3. Turn on the Nikon Optiphot Microscope and adjust the light to the desired brightness.
4. Use the 10× objective to find an empty area on the slide with no tissue. Use the Auto White button to set the white balance for the pictures.
5. Find the desired place to take a picture on the slide.
6. Turn the Nikon Digital Sight Camera to the left to switch the view from the microscope to the NIS-Elements BR software.
7. Use the fine focus knob on the microscope to bring the image into focus.
8. Open the Camera Settings and set Mode to Normal, Format Fast to 1,280 × 960, and Format Quality to 2,560 × 1,920. Leave all the other settings at their default settings.
9. Use the software's Auto Exposure button to find the optimal exposure and gain for the picture with the 10× objective. Maintain the same settings for all pictures taken with the 10× objective.
10. Ensure the image is focused using the fine focus knob on the microscope, then capture the image with the capture button.
11. Save the image as a tagged image file format (.tif).
12. Switch to the 40× objective and use Auto Exposure to find the optimal exposure and gain. Maintain the same settings for all pictures taken with the 40× objective.
13. Repeat the same process for capturing and saving the picture.
Data analysis
Data analysis procedures were described in our published research article [8]. The data analysis can be found in Figure 2 of the article [8]. Pten L/L mice were infected with Ad-CMV-Luc and Ad-Cre-Luc virus. Mice infected by Ad-CMV-Luc were used as Pten-positive control. The relative Pten expression level and the downstream signaling molecules, such as phosphorylated-AKT and phosphorylated-S6, were estimated using immunohistochemistry (IHC, procedures were described above). The number of recommended biological replications is ≥3. All sections (control and treated) must be stained at the same time using the same procedures. Sections with relatively higher protein levels develop darker color during this process.
Validation of protocol
The success of viral delivery is validated by the luciferase-based bioluminescent imaging results (Figure 2).
This protocol has been used and validated in the following research article:
• Liu et al. [8]. A Novel Controlled PTEN-Knockout Mouse Model for Prostate Cancer Study. Frontiers in Molecular Biosciences (Figure 2, Panels A–C).
In this article, the Pten conditional knockout mouse model was validated by estimating relative Pten expression levels in the prostate tissues using immunohistochemistry.
The figure from the original article is presented below (Figure 4).
Figure 4. Expression levels of PI3K/mTOR pathways' downstream components in mice's anterior prostate respond to Pten conditional knockout. This figure is adapted from the authors' published article, A Novel Controlled PTEN-Knockout Mouse Model for Prostate Cancer Study (Figure 2, Panels A–C). Frontiers in Molecular Biosciences [8]. (A) IHC staining to detect Cre recombinase in Pten LoxP/LoxP (L/L) mice's anterior prostate (AP) from different age groups that are infected with different adenovirus vectors. Ad-CMV-Luc, adenovirus vector that expresses luciferase using a cytomegalovirus (CMV) promoter. Green arrows indicate the basal cells. Green dash lines outline the stroma. Green triangles point toward stroma cells. Pten, phosphatase and tensin homolog deleted on chromosome 10. Ad-Cre-Luc, adenovirus vector that co-expresses luciferase and Cre recombinase (also using the CMV promoter). (B) IHC staining to detect Pten and downstream components of the PI3K/mTOR pathways in Pten L/L mice's AP that were infected with Ad-CMV-Luc and Ad-Cre-Luc viral vectors. Red arrows indicate Pten-, P-Akt-, P-S6-, or P-4E-BP1-positive cells. P-S6, phospho-S6 ribosomal protein. P-4E-BP1, phospho-eukaryotic translation initiation factor 4E-binding protein 1. (C) Serial IHC staining to detect Pten and downstream components of the PI3K/mTOR pathways in Pten L/L mice's AP at 0, 4, 8, and 16 weeks from the surgical delivery of the viral vectors. Red arrows indicate Pten-negative cells with activated P-S6 and P-4E-BP1.
General notes and troubleshooting
General notes
1. The rate of successful infection is approximately 80%. In other words, multiple mice with the same genotype must be prepared beforehand. In addition, always validate the viral delivery using luciferase-based bioluminescent imaging five days after the surgery.
2. The mouse may die in an anesthetic accident. Therefore, it is recommended that researchers work in pairs. The assistant can monitor the mouse's respiratory rate while the operator performs the surgery.
3. Viral infection may cause an immune response in mice. To eliminate this potential variable in cancer research, it is recommended to use mice with identical genotypes injected with a vacant viral vector as the control.
4. Mice purchased from outside usually have a heterozygous genotype (Pten LoxP/Wildtype, Pten L/WT). To generate homozygous (Pten L/L) mice, refer to the breeding strategy described in the supplementary information (Figure S1).
5. This model is designed for age-related cancer research. The supplementary material provides the timeframes of surgeries and examinations for different age groups (Figure S1).
Troubleshooting
Problems, critical steps, potential causes, and corresponding solutions are all listed below (Table 4).
Table 4. Troubleshooting
Problems Steps Causes Solution
No or little luminescent signals are detected B4 1. The mouse's immune system eliminated the virus. Perform the surgery on a different mouse.
2. Signal is weak due to the limited number of infected cells. Change the imaging parameters and scale of signals so that weak signals can be captured and displayed by the software.
3. Luciferase has not completely reacted with D-luciferin. Wait longer before image acquisition.
IHC overstaining G22–G36 Chromogenic reaction (incubation with DAB) takes too long. Start over with new sections. Shorten the chromogenic reaction (incubation with DAB) or use further diluted antibodies.
Insufficient IHC staining Chromogenic reaction occurs too quickly. Elongate the chromogenic reaction. If no more color is developed, start over with new sections and use more concentrated antibodies.
Hematoxylin overstaining G37–G44 Hematoxylin treatment takes too long. Extend the treatment with the ethanol-based 0.5% (v/v) HCl solution.
Insufficient hematoxylin staining Slides are soaked in the ethanol-based 0.5% HCl solution for too long. Repeat the hematoxylin staining.
Note: In the steps column, capital letters refer to Procedure subsections, and the numbers refer to the specific steps within the subsection denoted by the capital letter. For instance, B4 refers to step 4 in subsection B under Procedures. IHC, immunohistochemistry. DAB, 3,3′-diaminobenzidine. HCl, hydrogen chloride.
Supplementary information
The following supporting information can be downloaded here:
1. Figure S1. Breeding strategy and timeframe of surgeries and examinations for different age groups
Acknowledgments
This work was supported by the National Cancer Institute of the National Institutes of Health (NIH) under Award Number R01CA255802, the National Institute of General Medical Sciences of the NIH under Award Number P20GM103629, and U54GM104940, the Carol Lavin Bernick Faculty Grant and University Senate Committee on Research Fellowship Program Award from Tulane University. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health. Statistical analyses were completed in the Genomics and Biostatistics Core at the Tulane Center for Aging, which is supported by the National Institute of General Medical Sciences Grant P20GM103629.
The protocol was described and validated in this original research paper: Liu et al. [8]. A Novel Controlled PTEN-Knockout Mouse Model for Prostate Cancer Study. Frontiers in Molecular Biosciences.
Competing interests
The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.
Ethical considerations
The animal study was reviewed and approved by The Institutional Animal Care and Use Committee of Tulane University.
References
Brawer, MK. (2005). Prostatic intraepithelial neoplasia: an overview. Rev Urol. 7 Suppl 3: S11-8.
Freeland, J., Crowell, P. D., Giafaglione, J. M., Boutros, P. C. and Goldstein, A. S. (2021). Aging of the progenitor cells that initiate prostate cancer. Cancer Lett. 515: 28–35.
Sun, F., Báez-Díaz, C. and Sánchez-Margallo, F. M. (2017). Canine prostate models in preclinical studies of minimally invasive interventions: part I, canine prostate anatomy and prostate cancer models. Transl Androl Urol. 6(3): 538–546.
Koike, H., Nozawa, M., De Velasco, M. A., Kura, Y., Ando, N., Fukushima, E., Yamamoto, Y., Hatanaka, Y., Yoshikawa, K., Nishio, K., et al. (2015). Conditional PTEN-deficient Mice as a Prostate Cancer Chemoprevention Model. Asian Pac J Cancer Prev. 16(5): 1827–1831.
Leow, C. C., Wang, X. D. and Gao, W. Q. (2005). Novel method of generating prostate-specificCre-LoxP gene switching via intraductal delivery of adenovirus. Prostate. 65(1): 1–9.
Syyam, A., Nawaz, A., Ijaz, A., Sajjad, U., Fazil, A., Irfan, S., Muzaffar, A., Shahid, M., Idrees, M., Malik, K., et al. (2022). Adenovirus Vector System: Construction, History and Therapeutic Applications. Biotechniques. 73(6): 297–305.
Krakau, S., Richard, H. and Marsico, A. (2017). PureCLIP: capturing target-specific protein–RNA interaction footprints from single-nucleotide CLIP-seq data. Genome Biol. 18(1): 240.
Liu, S., Zhang, B., Rowan, B. G., Jazwinski, S. M., Abdel-Mageed, A. B., Steele, C., Wang, A. R., Sartor, O., Niu, T., Zhang, Q., et al. (2021). A Novel Controlled PTEN-Knockout Mouse Model for Prostate Cancer Study. Front Mol Biosci. 8: e696537.
Article Information
Publication history
Received: Jul 19, 2024
Accepted: Oct 23, 2024
Available online: Nov 11, 2024
Published: Jan 5, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Molecular Biology > DNA > Gene expression
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Cryopreservation Method for Preventing Freeze-Fracture of Small Muscle Samples
NG Namrata Ghag
JT Joshua Tam
RA R. Rox Anderson
NC Nashwa Cheema
Published: Vol 15, Iss 1, Jan 5, 2025
DOI: 10.21769/BioProtoc.5145 Views: 354
Reviewed by: Vivien J. Coulson-ThomasThirupugal Govindarajan Anonymous reviewer(s)
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Abstract
Histological techniques to study muscle are crucial for assessing skeletal muscle health. To preserve tissue morphology, samples are usually fixed in formaldehyde or cryopreserved immediately after excision from the body. Freezing samples in liquid nitrogen, using isopentane as a mediator for efficient cooling, preserves the tissue in its natural state. However, this method is highly susceptible to freeze-fracture artifacts, which alter or destroy tissue architecture. Isopentane is most commonly used in a semi-frozen/liquid state that is visually assessed by the experimenter, which can pose a challenge when freezing multiple tissues at a time or maintaining a consistent temperature. Furthermore, tissue size is also a confounding factor; depending on the size, freezing times can vary. In this study, we compare two different options for using isopentane while cryopreserving tissue. We also present an easy and reproducible method of freezing the soleus tissue of mice using frozen isopentane. This method decreased the occurrence of freeze-fractures by an order of magnitude, to ~4%, whereas the traditional method of cryopreservation resulted in ~56% freeze-fracturing.
Key features
• A uniform and highly reproducible protocol for freezing any tissue that is prone to freeze-fracture.
• Removes the need to maintain a mixed state of isopentane.
• Optimized cryopreservation method for the soleus muscle of mice.
• Allows for prevention of peripheral freeze-fracture in tissue, which is the most susceptible region to freeze-fracture damage.
Keywords: Freeze-fracture Soleus Muscle Mouse Cryopreservation Histology
Graphical overview
Background
Preserving tissue integrity is paramount for comprehensive morphological studies. Previous methods employed for tissue preservation, including formaldehyde fixation and cryopreservation, have demonstrated both advantages and challenges [1]. Formaldehyde fixation, while stabilizing tissues through protein crosslinking, may alter the natural conformation of cellular structures and impede antigen accessibility during immunohistochemical analysis. Cryopreservation, involving rapid freezing in liquid nitrogen without the need for fixation, is a widely used method for long-term tissue storage. However, the sudden temperature drop associated with liquid nitrogen often distorts the morphology of skeletal muscles. The liquid nitrogen evaporates immediately as it comes into contact with the warm tissue. This vapor becomes trapped between the mold surface and the liquid nitrogen, essentially creating an insulated barrier preventing freezing [2]. This process creates artifacts where intracellular water crystallizes immediately, resulting in the formation of huge crystals that rupture the muscle fibers. To address this, isopentane is utilized as a medium to facilitate controlled freezing [3].
A beaker filled with isopentane is placed in liquid nitrogen such that one-third of the isopentane is submerged. The isopentane is considered good for cryopreservation when a frozen layer forms at the bottom. At this stage, the bottom of the tissue mold is submerged in the chilled liquid. The duration of tissue submersion varies depending on the size of the tissue. Larger tissue requires longer freezing times than smaller tissues. The freezing times for small tissues like the diaphragm or extensor digitorum longus (EDL) is 6–12 s [3]. However, mice have even smaller tissues (e.g., soleus) that would require a freezing time of less than 6 s. Hence, small skeletal muscle tissues remain susceptible to freeze-fractures due to their higher surface-to-volume ratio. This method of freezing is highly inconsistent due to the many variables, i.e., liquid-to-frozen phase change, temperature fluctuations, visual assessment of phase state, tissue size, submerged depth of the mold, and batch-to-batch variability. In this study, rather than visual assessment, we monitor the temperature of isopentane when used for cryopreservation and present a reliable and reproducible method of preserving mouse skeletal muscles for both large and small tissues.
Materials and reagents
Biological materials
1. Tibialis anterior and soleus from 6-week-old C57BL/6 males (Jackson Laboratory, strain #000664)
Reagents
1. Bendable thermocouple probes for liquids and gases (McMaster Carr, catalog number: 39095K96)
2. 2-methylbutane/isopentane (Sigma-Aldrich, catalog number: 277258)
3. Dry ice
4. Liquid nitrogen
5. OCT (Optimal Cutting Temperature) (Sakura, catalog number: 4583)
6. CitriSolv (Fisher Scientific, catalog number: 04-355-121)
7. Bluing reagent (Fisher Scientific, catalog number: 22-050-114, Supplier No.: 7301)
8. Formalin (Fisher Scientific, catalog number: SF100-20)
9. Ethanol (Decon Laboratories, Inc, catalog number: 2701)
10. Hydrochloric acid (HCl) (Fisher Scientific, catalog number: A144-500)
11. Hematoxylin 560-Surgipath (Leica Biosystems, catalog number: 3801570)
12. Alcoholic eosin Y515-Suripath (Leica Biosystems, catalog number: 3801615)
Laboratory supplies
1. Modified big aluminum can (23Fl Oz), see section B: Cryopreservation method #1
2. Modified small aluminum can (11.5Fl Oz), see section C: Cryopreservation method #2
3. Handheld thermometer (McMaster Carr, catalog number: 9281T52)
4. Styrofoam ice box
5. Tissue molds (VWR, catalog number: 25608-916, Supplier No.: 4557)
6. Needles 23G (Exel International, catalog number: 26407)
7. Microscope slides (Epic Scientific, catalog number: 187200810RE)
8. Superfrost plus slides (Fisher Scientific, catalog number: 1255015)
Equipment
1. Blunt and sharp forceps (Fisher Scientific, catalog numbers: 12740916 and 11340705)
2. Scissors (Fisher Scientific, catalog number: 08-940)
3. Locking forceps (Fisher Scientific, catalog number: 13-812-16)
4. Cryostat blade (C.L. Sturkey, Inc. catalog number: 22-210-045)
5. Nanozoomer (Hamamatsu, model: C9600-12, catalog number: 000382)
Software and datasets
1. NDPI viewer software (Hamamatsu version 2.9.29)
Procedure
A. Dissection of muscle
1. Isolate tibialis anterior (TA) and soleus muscles from mice (6-week-old C57BL/6 males were used here) as previously described [4].
2. Cut each muscle in half at the mid-belly and place each half in molds filled with OCT. Use a needle to orient the muscle so that the cross-sectional side faces the bottom of the mold (Figure 1A).
Figure 1. Muscle orientation and equipment used for cryopreservation. A, B) TA and soleus were cut at the mid-belly and placed in molds containing OCT. The tissue was placed at the bottom half of the mold; A) top view and B) side view. C) The instrument setup for cryopreservation methods was laid out on the bench. Temperature probe and temperature meter were used to monitor the temperatures in both methods. Isopentane was filled up to the marks in the aluminum can. OCT was used to embed the tissues. Forceps were used to insert the molds in the cans with isopentane. A styrofoam box with dry ice was kept ready to temporarily hold the molds until they were transferred to a -80 °C freezer. Not shown in the picture are the two differently sized styrofoam boxes (large for larger can/small for smaller can) containing liquid nitrogen.
3. Confirm orientation visually from the side (Figure 1B). By using this technique for embedding, the experimenter already has an understanding of the tissue location and orientation. This method also prevents the tissue from easily floating in the OCT while the mold is being handled by the experimenter.
B. Cryopreservation method #1
This is the conventional method for freezing muscle tissues for histology [1].
1. Cut the front part of the large aluminum can with a pair of scissors as shown in Figure 1C. This will allow the evaporation of isopentane fumes resulting from the contact of room-temperature mold with the colder isopentane. This way, the experimenter can easily visualize the mold when placed inside the can.
2. Fill the can with approximately 300 mL of isopentane and place it in a styrofoam box containing liquid nitrogen such that one-third of the can, containing 100 mL of isopentane, is submerged. Mark the level of isopentane inside the can with a sharpie (Figure 1C).
3. Use a larger can for cryopreserving in liquid isopentane as more volume is required.
4. To easily transfer an open-filled can, keep one side of the can taller to make it easier to hold. Attach a paper towel and binder clip to the can to protect the user from the cold temperature when handling the can.
5. Freeze the isopentane until the bottom part of the can has a frozen white layer and temperature reaches -120 °C (Figure 2). Place the temperature probe on the top of the isopentane layer where the mold will be placed.
Figure 2. Detailed process of preserving the muscle tissue by two different methods. A) A bigger aluminum can was filled up to the mark with isopentane and inserted in a styrofoam box filled with liquid nitrogen. The temperature of the liquid isopentane was checked after the bottom layer was frozen (changes from translucent to white). Once the temperature reached -120 °C, the tissue mold was held in the liquid isopentane with forceps and freezing of the tissue was achieved. Precaution was taken not to submerge the tissue mold inside the isopentane. It was transferred with forceps to dry ice until storage at -80 °C. B) A small aluminum can was filled up to the mark with isopentane. It was placed inside a small styrofoam box filled up to one-third with liquid nitrogen to achieve quick and complete freezing of isopentane. Frozen isopentane was allowed to reach a temperature of -160 °C (as shown in the temperature meter); the tissue mold was placed on the frozen surface. Tissue mold was transferred using forceps on dry ice until storage in a -80 °C freezer.
6. Prepare the mold with tissue embedded in OCT. Hold the mold with locking forceps and place it in the chilled isopentane, ensuring the mold edges rest on the serrations to prevent tilting (Video 1).
Video 1. Cryopreservation method 1 and 2
7. Do not submerge the mold completely.
8. Freeze until OCT turns completely white. The outer border begins to freeze first and expands to the center of the mold. When the entire OCT turns from colorless to white, transfer the frozen mold to dry ice for 5–10 min to allow any residual isopentane to evaporate.
9. Store tissue samples at -80 °C until further analysis.
C. Cryopreservation method #2
1. Trim a small aluminum can and fill it with approximately 30 mL of isopentane to completely cover the bottom and maintain a flat, leveled surface on the top.
2. Freeze the isopentane completely by submerging the can in a styrofoam box filled to one-third with liquid nitrogen. As smaller volumes are needed when isopentane is used in the solid state, no added support is required. The same paper towel and binder clip can be used with the small can to allow the user to handle it safely.
3. Ensure the temperature of frozen isopentane reaches -160 °C (freezing point of isopentane) by placing the tip of the temperature probe on it (Figure 2).
4. Prepare the mold with tissue embedded in OCT. Place it on the frozen surface of isopentane using locking forceps (Video 1). Holding the mold with forceps is unnecessary, as the isopentane surface is solid.
5. Freeze until OCT turns completely white. Using forceps, transfer the frozen mold to dry ice for 15–20 min to allow residual isopentane to evaporate.
6. Wrap the mold in aluminum foil or place it in a sample bag.
7. Store tissue samples at -80 °C until further use.
D. Cryosectioning
1. Remove tissue samples from the -80 °C freezer and allow them to equilibrate to -20 °C in the cryostat.
2. Place tissue blocks on the cryostat sample holder with the cross-sectional side of the tissue facing up.
3. Cut the tissue blocks to produce 10 µm thick sections.
4. Place sections on Superfrost plus slides and store the slides in a -80 °C freezer.
E. H&E staining
1. Thaw slides to room temperature and air-dry them for 30 min.
2. Fix slides in formalin for 1 min, then wash them in distilled water for 2 min.
3. Dip slides in hematoxylin stain for 3 min. Hematoxylin stains the nuclei and gives it a reddish-purple color.
4. Wash slides in distilled water for 2 min.
5. Briefly dip slides in 0.25% acid alcohol (made from ethanol and HCl) to remove excess stain and then wash them in distilled water for 2 min.
6. Dip slides in bluing reagent for 1 min to change the previously stained nuclei to a blue color and then wash them in distilled water for 2 min.
7. Immerse slides in 95% ethanol for 30 s, followed by eosin stain for 1.5 min.
8. Wash slides in 95% ethanol for 30 s.
9. Dip the slides for 1 min in three separate containers with absolute ethanol.
10. Wash slides for 3 min in three separate containers with CitriSolv reagent.
11. After staining, image the slides using a Nanozoomer for analysis. H&E images for TA and soleus are presented in Figures 3 and 4, respectively.
Figure 3. Comparison of H&E images of mouse tibialis anterior (TA) muscles embedded via liquid and frozen isopentane. A) Three TA muscle tissues were used for each of the preservation methods. Cell cytoplasms appear pink, and nuclei are stained blue. Images were zoomed in at randomly selected areas to look for freeze-fractures. Freeze-fracture was evident in tissues preserved by liquid isopentane. B) TA muscle preserved using frozen isopentane shows the absence of freeze-fractures and displays increased tissue integrity. Scale bar: 100 μm.
Figure 4. Comparison of H&E images of mouse soleus muscles embedded via liquid and frozen isopentane. A) Three soleus muscles were used for each of the preservation methods. As soleus is smaller than tibialis anterior (TA) and most of the other mouse muscles, it is more prone to freeze-fracture. Images were zoomed in to look for freeze-fractures. Freeze-fracture was evident in tissues preserved by liquid isopentane. B) Soleus muscles preserved using frozen isopentane show the absence of freeze-fractures. Scale bar: 100 μm.
Data analysis
1. Five soleus muscle tissues were selected per method.
2. Total tissue area was determined by outlining the tissue in NDPI viewer software.
3. Multiple regions of damage were identified and outlined to determine their area. All the areas were summed to obtain a total freeze-fracture area.
4. Percentage of freeze-fracture was calculated. A mean of freeze-fracture percentage was obtained.
5. Statistical comparison was performed using the Student’s t-test. A p-value equal to or less than 0.05 was considered statistically significant.
Validation of protocol
Mouse muscle tissues are often frozen using ice-cold isopentane in liquid state to minimize tissue damage. This method is highly susceptible to variations caused by the phase state of isopentane and the duration of the technique, resulting in freeze-fracture in tissues. We optimized this method of cryopreservation using frozen isopentane. We validated this protocol by comparing the extent of freeze-fracture in both methods using soleus muscle and observed that when using frozen isopentane, tissue damage is minimal (~4%) compared with when freezing with liquid isopentane (~56%) (Table 1). We observed a significant difference between the two methods (N5/method). Additionally, when using frozen isopentane, the entire tissue morphology is protected. The periphery of the tissue is highly susceptible to freeze-fracture when cryopreserved in liquid isopentane; however, with the use of frozen isopentane, freeze-fracture is minimal at the periphery (Figure 5).
Table 1. Measurement of freeze-fractured area in H&E
Total area (mm2) Freeze-fracture area (mm2) % Freeze-fracture
Liquid Frozen Liquid Frozen Liquid Frozen
1.3 1.8 0.5 0.14 40 8
1.4 1.4 1.15 0.04 81 3
0.8 1.7 0.29 0.05 36.1 2.8
1.5 1.5 1.29 0.06 88.9 4.1
1 1.2 0.33 0.02 34.2 1.5
Mean 56.04 3.88**
SD 26.62073 2.48133
** p < 0.01 between liquid vs. frozen.
Figure 5. Identifying regions of freeze-fracture in soleus muscles. A) The tissue boundary is most susceptible to damage by freeze-fracture compared with the center. In the liquid isopentane method, tissue periphery displays significant freeze-fracture, as indicated by the greyed-out area. Zoomed views of the peripheral vs. central regions of the tissue. B) The frozen isopentane method is critical in eliminating this peripheral freeze-fracture to the tissue.
General notes and troubleshooting
1. A cold-resistant PTFE beaker or a cryogenic bowl/dish can be used instead of a modified aluminum can.
2. A large styrofoam ice box was used for the larger can. Liquid nitrogen was added to the box such that one-third of the can was submerged. The can was placed in the corner of the box for added support and to prevent the can from tipping over in the ice box. A smaller styrofoam ice box was used for the small can. The can occupied one corner of the box and was the appropriate size to be securely placed. Liquid nitrogen was added such that one-third of the can was submerged. As the box was smaller, very little liquid nitrogen was needed.
3. As the room-temperature mold containing the sample is placed on the frozen isopentane, fumes begin to form until the mold reaches the temperature of the isopentane. Meanwhile, the frozen surface beneath the mold will melt. If the mold is left too long and the can is still in liquid nitrogen, eventually the liquid isopentane will refreeze, and the mold may become stuck. In that case, remove the can and quickly extract the mold as soon as it becomes loose.
Acknowledgments
This work was funded in part by the Musculoskeletal Injury Rehabilitation Research for Operational Readiness (MIRROR), Department of Physical Medicine & Rehabilitation, Uniformed Services University, Bethesda, MD, and The Wellman Center, Boston, MA collaboration (HU00011920056). The views expressed herein are those of the authors and do not necessarily reflect the official policy or position of the Uniformed Services University, Defense Health Agency, Department of Defense, or the U.S. Government. R.R.A. was partially supported by the Lancer Endowed Chair in Dermatology.
Competing interests
The authors confirm that there are no known financial interests or personal relationships that could have influenced the work reported in this paper.
Ethical considerations
The authors state that there are no ethical conflicts associated with this research.
References
Leiva-Cepas, F., Ruz-Caracuel, I., Peña-Toledo, M. A., Agüera-Vega, A., Jimena, I., Luque, E. and Peña-Amaro, J. (2018). Laboratory methodology for the histological study of skeletal muscle. Arch Med Deporte. 35: 254–262.
Dubowitz, V., Sewry, C. and Oldfors, A. (2021). Preface to the Fifth Edition. In: Dubowitz, V., Sewry, C. A. and Oldfors, A. (Eds.). Muscle Biopsy (Fifth Edition). pp. vii. Elsevier.
Kumar, A., Accorsi, A., Rhee, Y. and Girgenrath, M. (2015). Do's and Don'ts in the Preparation of Muscle Cryosections for Histological Analysis. J Visualized Exp. (99): e52793.
Wang, C., Yue, F. and Kuang, S. (2017). Muscle Histology Characterization Using H&E Staining and Muscle Fiber Type Classification Using Immunofluorescence Staining. Bio Protoc. 7(10): e2279.
Article Information
Publication history
Received: Aug 1, 2024
Accepted: Oct 18, 2024
Available online: Nov 12, 2024
Published: Jan 5, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Cell Biology > Cell isolation and culture > Cryopreservation
Cell Biology > Tissue analysis > Histomorphology
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5,146 | https://bio-protocol.org/en/bpdetail?id=5146&type=0 | # Bio-Protocol Content
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A Novel and Robust Method for Investigating Fungal Biofilm
BB Biswambhar Biswas
SA Shumaiza Asif
RP Rekha Puria
AT Anil Thakur
Published: Vol 15, Iss 1, Jan 5, 2025
DOI: 10.21769/BioProtoc.5146 Views: 474
Reviewed by: Lucy XieSascha BrunkeSimab KanwalShailesh Kumar
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Original Research Article:
The authors used this protocol in Microbiology Spectrum Aug 2023
Abstract
Candida auris, labeled an urgent threat by the CDC, shows significant resilience to treatments and disinfectants via biofilm formation, complicating treatment/disease management. The inconsistencies in biofilm architecture observed across studies hinder the understanding of its role in pathogenesis. Our novel in vitro technique cultivates C. auris biofilms on gelatin-coated coverslips, reliably producing multilayer biofilms with extracellular polymeric substances (EPS). This method, applicable to other Candida species like C. glabrata and C. albicans, is cost-effective and mimics the niche of biofilm formation. It is suitable for high-throughput drug screening and repurposing efforts, aiding in the development of new therapeutics. Our technique represents a significant advancement in Candida biofilm research, addressing the need for consistent, reproducible biofilm models. We detail a step-by-step procedure for creating a substratum for biofilm growth and measuring biofilm thickness using confocal laser scanning microscopy (CLSM) and ultrastructure by scanning electron microscopy (SEM). This method provides consistent outcomes across various Candida species.
Key features
• The biofilm formed on gelatin surfaces mimics host conditions, replicating the multilayered structure and EPS, offering a more accurate model for studying C. auris biofilms.
• This method is highly reproducible and suitable for drug screening and biofilm analysis through three-dimensional (3D) reconstruction.
• This in vitro technique aids in studying biofilm formation, related virulence properties, and drug tolerance of C. auris and other Candida species.
• The simple, cost-effective technique is ideal for screening novel inhibitors and repurposed drug libraries, facilitating the design/identification of new therapeutics against Candida species.
Keywords: Candida albicans Candida auris Candida glabrata EPS Gelatin Biofilms
Graphical overview
Background
Candidiasis is a serious threat, especially to individuals with compromised immune systems and comorbidities. Hospitalized patients, particularly those who have undergone organ or bone marrow transplants or are receiving immune-suppressive therapies, are highly susceptible to infections [1]. Prolonged hospital stays further increase the risk of contracting hospital-acquired infections [2]. A critical concern is the emergence of Candida auris, a fungal pathogen first isolated in 2009 [3]. Since then, C. auris has become a global health concern, with reports of drug resistance emerging from as many as 32 countries, particularly during the COVID-19 pandemic among hospitalized patients [4]. C. auris as a drug-resistant nosocomial pathogen has a reported mortality rate of 30%–50% in infected persons [5]. This pathogen can cause various infections, including ear, systemic (fungemia), wound, lung, urinary tract, and gut infections [5–7]. Due to its high resistance to various antifungal treatments and its potential for pan-drug resistance, the World Health Organization (WHO) has listed C. auris in the critical priority group of fungal pathogen priority list (FPPL) released in October 2022 [8].
Unlike other Candida species, C. auris seldom colonizes the gut but can persist on the skin by forming resilient biofilms, which causes rapid spread of this fungus in hospitals [9]. C. auris demonstrates remarkable resilience, especially in its biofilm form, which shows substantial resistance [10]. According to the Centers for Disease Control and Prevention (CDC), C. auris biofilms are resistant to common disinfectants, including ammonium salt–based solutions, and can endure harsh conditions [11]. C. auris can persist in the cutaneous layer by dividing and forming biofilms, which can cause bloodstream infection when they reach the blood vessels [12]. This underscores the necessity of studying biofilm formation in infection scenarios to develop effective treatments. Given the widespread drug resistance, there is an urgent need for robust screening platforms to identify novel inhibitors. Drug repurposing must be validated through rigorous in vitro and in vivo screening methods. However, a major challenge is establishing a reliable method to form biofilm models that accurately replicate in vivo conditions. The phenotypes of C. auris biofilms can vary significantly based on culture conditions, complicating research outcomes. Current methods, such as those using Thermanox (polystyrene) coverslips or porcine skin surfaces [13,14], either fail to produce typical biofilm architecture or lack cost-effectiveness for large-scale screenings. A standard surface for biofilm proliferation is crucial for reproducibility, as effective drug screening requires the typical multilayered architecture of biofilms.
To address these issues, we proposed a novel in vitro method for growing C. auris biofilms on gelatin-coated coverslips derived from bovine skin. Gelatin, a derivative of collagen, mimics the matrix of mammalian cells, thereby more closely replicating natural conditions. This method successfully forms heterogeneous, multilayered biofilms of C. auris. The architecture of these biofilms has been analyzed using scanning electron microscopy (SEM) and compared with previous studies [14–16]. Furthermore, the versatility and accuracy of this method have been demonstrated using confocal scanning laser microscopy (CSLM) to measure biofilm thickness. The application of this innovative in vitro technique is not only limited to C. auris biofilms but can also be used to study biofilm formation in other Candida species, such as Candida glabrata and Candida albicans. This method is a potent, cost-effective solution for large-scale drug screenings.
Here, we describe a stepwise procedure for creating a substratum for biofilm growth and measuring biofilm thickness with confocal laser scanning microscopy and ultrastructures by scanning electron microscopy. As mentioned above, it results in reliable outcomes and is suitable for various Candida species with different media. This method has been successfully used in previously published studies, demonstrating its effectiveness and reliability [15,16].
Materials and reagents
Biological materials
1. Candida auris strain of clade 2, CBS10913T
Reagents
1. Gelatin derived from bovine skin and bones (Sigma-Aldrich, catalog number: G9391-100G)
2. Double-distilled water
3. BD Difco YPD broth (Fisher Scientific, catalog number: DF0428-17-5)
4. 0.3 N hydrochloric acid (HCl) (Fisher Scientific, catalog number: 7647-01-0)
5. 100% ethanol (G-Bioscience, catalog number: RC1533)
6. 3-aminopropyl triethoxysilane (Himedia, catalog number: RM6592-100G)
7. Acetone HPLC grade (MERCK, catalog number: 60002010001730)
8. Glutaraldehyde 25%, w/w (Himedia, catalog number: MB222-100ML)
9. RPMI-1640 w/ or w/o MOPS and with L-glutamate added (Himedia, catalog number: AL028G-500ML)
10. Calcofluor white (CFW) (Sigma-Aldrich, catalog number: 18909-100ML-F)
11. Paraformaldehyde (Sigma-Aldrich, catalog number: F8775-500ML)
12. Alcian blue (Himedia, catalog number: RM471-25G)
13. Sodium cacodylate trihydrate (Himedia, catalog number: RM3732-25G)
14. Sodium chloride (NaCl) (Himedia, catalog number: MB023-1KG)
15. Potassium chloride (KCl) (G Biosciences, catalog number: RC1167)
16. Sodium phosphate dibasic anhydrous (Na2HPO4) (SRL, catalog number: 53046)
17. Potassium phosphate monobasic (KH2PO4) (G Biosciences, catalog number: RC1173)
18. Transparent nail paint
Solutions
1. YPD broth (see Recipes)
2. 1× PBS (see Recipes)
3. Scanning electron microscopy (SEM) buffer (see Recipes)
4. Other solutions (see Recipes)
Recipes
1. YPD broth (1 L)
Add 50 g of Difco YPD broth to 1 L of distilled water and sterilize by autoclaving at 121 °C for 20 min before use.
2. 1× PBS (1 L)
To prepare 1× PBS, weigh the components mentioned in the table below and dissolve in 500 mL of water. Once all the components are dissolved, make up the volume to 1 L and autoclave.
Reagent Amount
NaCl 8 g
KCl 0.2 g
Na2HPO4 1.44 g
KH2PO4 0.24 g
Double-distilled water (ddH2O) to 1 L
3. Scanning electron microscopy (SEM) buffer (50 mL)
To prepare the SEM buffer, add the following chemicals and make up the volume to 50 mL. Filter with pore size 0.22 μm, sterilize, and store at -20 °C.
Reagent Stock concentration Amount Final concentration
Alcian blue n/a 75 mg 0.15%
Sodium cacodylate n/a 1.2 g 0.15 M
Glutaraldehyde 25% 4 mL 2%
Paraformaldehyde 37% 2.7 mL 2%
Double distilled water (ddH2O) n/a To 50 mL n/a
Total 50 mL
4. Other solutions
Reagents Stock concentration Amount Final concentration
CFW 1 mg/mL 30 μL 30 μg/mL in 1 mL
HCl 12 N 2.5 mL 0.3 N in 100 mL
3-aminopropyl triethoxysilane n/a 1 mL 2% in 50 mL
Gelatin n/a 1g 1% in 100 mL
Ethanol 100% 2.5 mL (10 mL) 25%
100% 5 mL (10 mL) 50%
100% 7 mL (10 mL) 70%
100% 8 mL (10 mL) 80%
100% 9.5 mL (10 mL) 95%
Laboratory supplies
1. 1.5 mL microcentrifuge tubes (Tarsons, catalog number: 500010)
2. 15 mL conical bottom tubes (Tarsons, catalog number: 430766)
3. 50 mL conical bottom tubes (Tarsons, catalog number: 430829)
4. 1,000 μL tips (Tarsons, catalog number: 521020B)
5. 200 μL tips (Tarsons, catalog number: 521010Y)
6. Conical flasks (Borosil catalog number: 4980021)
7. 90 mm Petri dishes (Tarsons, catalog number: 460095)
8. 24-well polystyrene plates (Thermo Scientific, catalog number: 142475)
9. 6-well polystyrene plates (Thermo Scientific, catalog number: 140675)
10. Kimwipes (Fisher Scientific, catalog number: 34120)
11. Glass slides (any high-quality glass slides)
12. Glass coverslips (any high-quality glass coverslips)
13. Carbon tape (TED PELLA, Inc., product no.: 16084-1)
Equipment
1. Eppendorf New Brunswick InnovaTM 42/42 R, stackable incubator shaker (Fisher Scientific, catalog number: 05-400-162)
2. Heat block (DTH-100 Dry bath incubator)
3. Vortex (BR Biochem, catalog number: BI-VM-2500)
4. Refrigerated benchtop centrifuge 5910 R (Eppendorf, catalog number: 5942000130)
5. Centrifuge 5427 R (Eppendorf, catalog number: 5429000133)
6. Light microscope (Leica, model: Sp5)
7. Scanning electron microscope (Apreo VolumeScope FEI)
8. Airstream® class II type A2 biological safety cabinet
9. Spectrophotometer Gene Quant 1300 (VWR, catalog number: SCLI80-2120-02)
10. Vacuum desiccator (Sigma-Aldrich, catalog number: Z119024)
11. Quorum SC7620 sputter coater
12. Hot-air oven (Scientific Systems)
13. Digital orbital shaker (Heathrow Scientific, catalog number: 120460)
14. Forceps
15. Autoclave
16. Water bath (Grant instruments, model: SBB Aqua 5 plus)
Software and datasets
1. LAS-AF
2. FIJI 1.54
3. GraphPad Prism 9
4. Adobe Photoshop CC 2019
Procedure
A. Preparation of coverslips
1. Gelatin preparation
a. Dissolve 1 g of gelatin (bovine skin derived) in 25 mL of ice-cold distilled water.
b. Vortex the solution to remove any clumps.
c. Centrifuge the solution at 1,000× g for 10 min at 4 °C.
d. The pellet consists of gelatin, while the top water layer contains debris, dust specks, and other water-soluble impurities.
e. Repeat this step ~10 times or until the top water layer is transparent.
f. Finally, resuspend the washed gelatin in 100 mL of water to make 1% w/v suspension and autoclave.
g. Store the autoclaved gelatin at -20 °C in 50 mL conical bottom tubes. This can be stored for up to 6 months to a year.
h. To warm the gelatin again, incubate the gelatin-containing tubes in a 37 °C water bath.
2. Coverslip preparation
a. All steps must be done in a sterile environment.
b. Treat coverslips in a conical flask half filled with 0.3 N hydrochloric acid and frequently swirl overnight using an orbital shaker at 100 RPM to dissolve any contaminations and organic impurities.
c. Remove the acid and thoroughly wash the coverslips with 50 mL of autoclaved water three times.
d. Treat the coverslip with 100% ethanol for 30 min, with frequent swirling.
e. Discard the ethanol and dry the coverslips in a hot-air oven at 65 °C overnight in a sterile flask.
f. Prepare a 2% (3-aminopropyl) triethoxysilane solution in acetone. Dip each coverslip in the silane solution two times using forceps. This alkoxysilane treatment helps bridge the organic groups of gelatin to the silicon on the glass surface.
g. Dry the coverslips at room temperature and then wash them with water to remove excess unbound silane.
h. Dry the coverslips thoroughly in a hot air oven for 6–8 h.
i. Dip the alkoxysilane-treated coverslips in the previously prepared warmed gelatin solution to coat the surfaces.
j. Dry the coverslips at room temperature for 15 min.
k. Dip the coverslips one by one in glutaraldehyde solution in a 50 mL conical bottomed tube using forceps and leave them to dry at room temperature.
l. Repeat the gelatin coating and glutaraldehyde fixation step again.
m. After the coating dries up, dip the coverslips in water for washing and drying.
n. These coverslips can be stored at -20 °C for a year in Petri dishes between lint-free tissue stacks.
B. Biofilm formation and processing
1. Biofilm formation
a. Inoculate fungus cells in 5 mL of YPD medium in a 50 mL flask or 50 mL tube and incubate at 30 °C and 200 RPM overnight to establish a primary culture.
b. Use overnight-grown culture to prepare a secondary culture at ≤ 0.1 OD600 and incubate at 200 RPM in a shaker incubator at 30 °C until it reaches an OD600 between 0.6 and 0.8. Measure OD600 in a spectrophotometer.
c. Harvest cells by centrifugation at 3,500× g for 5 min, wash the cell pellet twice with 1× PBS, and resuspend in 1 mL of 1× PBS.
d. Prepare RPMI 1640 or RPMI-MOPS medium for biofilm formation. RPMI 1640 or RPMI-MOPS can be used to grow the biofilm; the only difference is in the buffering systems present in both media. RPMI 1640 contains sodium bicarbonate as a buffer to maintain the pH, whereas RPMI-MOPS [3-(N-morpholino) propanesulfonic acid] contains propane sultone buffer with a pH of 7.2, maintaining the physiological buffering conditions.
e. Place individually coated coverslips into the wells of 24-well plates and add 500 μL of the appropriate medium to each well.
f. Adjust the cell density to an OD600 of 0.5 before seeding them onto the coverslips.
g. Adjust 0.5 OD600 cells over the coverslips. Allow the cells to adhere by incubating them for 2.5–3 h in an incubator maintained at 37 °C.
h. When changing the medium, avoid disturbing adhered cells by gently adding the medium in drops along the wells’ walls. This marks the starting point (Time point 0) and any treatment regime can be designed relative to this time point.
i. Incubate the biofilm for 48 h undisturbed at 37 °C in a static incubator.
j. After incubation, harvest the coverslips for further processing. To harvest the biofilm, carefully remove the coverslips from the wells using forceps without shaking or disturbing them to prevent disrupting the formed biofilm.
2. Biofilm processing for confocal laser scanning microscopy (CLSM)
a. Wash the biofilm-containing coverslips with 500 μL of 1× PBS buffer by slowly aspiring the media from the wells by its meniscus using a 200 μL pipette, followed by adding 1× PBS in a continuous dropwise manner along the walls.
b. The biofilm forms on the upper side of the coverslips. The calcofluor-white (CFW) signal will be calculated from the bottommost signal layer to the topmost layer to measure the biofilm thickness.
c. Prepare calcofluor-white (CFW) stain solution at a working concentration of 30 μg/mL dissolved in 1× PBS.
d. Add enough CFW to submerge the biofilm-containing coverslips in the wells and incubate in the CFW solution at 30 °C for 30 min.
e. Transfer the coverslips onto glass slides in such a way that the side with the biofilm faces the glass.
f. Seal the coverslips over the glass slides with transparent nail paint to prevent drying before visualization under the confocal microscope.
3. Biofilm processing for scanning electron microscopy (SEM)
a. Treat the biofilm-containing coverslips with 0.15 M sodium cacodylate by submerging them in this solution for 5 min at room temperature in 24- or 6-well plates and then discarding the remaining liquid. If using 24-well plates, 500 μL of solution is required to fully submerge the coverslip. For 6-well plates, approximately 3 mL of solution is needed to immerse the coverslips. Repeat this step two times.
b. Keep the biofilm-containing coverslips in SEM buffer (2% paraformaldehyde, 2% glutaraldehyde, 0.15 M sodium cacodylate, and 0.15% alcian blue in water) and incubate overnight at 4 °C.
c. Transfer the coverslips to another 6-well plate and wash them twice with 0.15 M sodium cacodylate.
d. Different concentrations of ethanol, such as 25%, 50%, 80%, and 95%, are made using absolute ethanol and 0.15 M sodium cacodylate in separate conical-bottom tubes.
e. Slowly dehydrate the fixed biofilm-containing coverslips by sequentially exposing them to increasing concentrations of ethanol in gradient, gradually starting with 25% for 5 min followed by 50% for 10 min, 70% for 15 min, 80% for 15 min, 95% for 30 min, and at last with absolute ethanol for 1 h.
f. Vacuum dry the biofilm-containing coverslips overnight in a vacuum desiccator.
g. Dry the coverslips by submerging them in acetone for 3 h.
h. After dehydration, mount the coverslips on adhesive carbon tape with the biofilm side facing up.
i. Coat the coverslips with gold particles using an argon ion beam coater, maintaining a pressure of 0.08 mbar and voltage of 15 mV.
C. Acquisition of images
1. Image acquisition using Leica SP5
a. Configure LAS-AF software with a pixel size set to 2,048 × 2,048 and a scanning speed of 200 Hz.
b. Utilize a 20× objective lens for image capture.
c. Select focus planes ranging from the lowest focal plane displaying blue signals to the upper focal plane showing blue signals. Blue signals depict the CFW bound to chitin.
d. Set step sizes at 0.6 μm for Z-stacking.
e. Perform scanning starting from the nearest dark slice in the lowest layer to the nearest dark slice in the highest layer.
f. Use bidirectional scanning mode and maintain a pinhole size of 60 μm so that a single layer comes at each step (Figure 1A).
Figure 1. Analysis of biofilms using confocal laser scanning microscopy. A) Single cross-section of the biofilms. All such cross-sections were captured along the Z-axis, resulting in a 3-dimensional reconstruction. B) From 1A, the thickness of the biofilm was measured. C) First, the image was opened in FIJI, and from the Bio-Formats Import options, “hyperstack” and “composite” were chosen. D) The desired field to be analyzed was selected again. E) The image was opened as shown. The default scale was applied by the LasAF imaging software during the acquisition. From the FIJI main menu, the set scale option from the Analyze tab was selected. F) The set scale tab appeared, and scale was noted down. G) From the Plugins tab, the 3D-Viewer was selected. H) The image was opened in the ImageJ 3D viewer module. I) It was set in the side view position. J) From the view tab in the ImageJ 3D viewer module, a snapshot of the image was taken. K) A snapshot was saved using the default image resolution. L) Then, the previous scale values from the set scale option under the Analyze tab were used. M) The image was converted into an 8-bit image. N) The threshold was set to subtract the background from the signal. O) The image was then ready for measurement. P) The line tool from FIJI tools was used to measure thickness. Q–R) The thickness of the biofilm was measured by pressing Ctrl + M. S) This shows the measured value of the biofilm thickness.
2. Image acquisition in Apreo Volume Scope FEI
a. Set the voltage to 10 kV and image at different magnifications (250×, 1,250×, 2,500×, and 5,000×).
b. An image captured at 2,500× is shown in Figure 2A.
D. Analysis of images
1. Drag and drop the captured images onto FIJI.
2. Select the color mode to composite and view stack with hyperstack (Figure 1C–E) in the Bio-Formats Import options.
3. Export a representative image of the top view in JPEG or TIFF format by navigating to File > Save As > TIFF or JPEG.
4. Reopen the TIFF or JPEG image in FIJI by dragging and dropping it onto the software.
5. Adjust the brightness and contrast by going to Image > Adjust > Brightness/Contrast and modifying the bars, then click Apply.
6. For the original imported image, set the scale either by using the imaging software’s preset or by going to Analyze > Set Scale and specifying the parameters (Figure 1F).
7. Draw the scale bar to the desired length using the Straight tool in the FIJI toolbox.
8. Create the scale bar by navigating to Analyze > Tools > Scale Bar.
9. Open the 3D viewer by going to Plugins > 3D Viewer (Figure 1G).
10. In the pop-up add window, select only the blue channel.
11. Set the resampling factor to 2 and click OK. Wait for buffering to complete.
12. Now, adjust the position using the mouse to ensure the thickness is visible and zoom in until the two side red borders meet the edge (Figure 1B and I).
13. From the ImageJ 3D Viewer view option, select Take snapshot and choose the original image’s resolution (e.g. 2,048 × 2,048) (Figure 1J–K).
14. Set the scale again by navigating to Analyze > Set Scale and manually entering the same parameters as in the original image (Figure 1L).
15. Convert the image to an 8-bit image by going to Image > Type > 8-bit (Figure 1M).
16. Set the threshold by going to Image > Adjust > Threshold, keeping parameters at Default and B&W (Figure 1N).
17. Adjust the second slide bar to make the signal prominent from the background and close the dialogue.
18. Select the Straight tool from the toolset and draw a straight line perpendicular to the planes to cover the thickness (Figure 1P–R).
19. Press Ctrl + M to measure the biofilm thickness. Take at least 25 measurements across the biofilm from each image.
20. A results tab will appear containing all the measurements. Use only the length of information and record it in a table format, ensuring the angle is ~90° (Figure 1S).
21. Plot the data in GraphPad Prism to create graphs and test the significance between replicates and test conditions.
E. Pseudocoloring: for better visualization and differentiation
1. Perform pseudocoloring in Adobe Photoshop CC 2019 using overlay mode (Figure 2A and B).
2. Select different objects seen using the quick selection tool and create a new layer.
3. Use the Paint Bucket tool to fill in color in the selection in the new layer.
4. Set the blending mode of the new layer to multiply.
Figure 2. SEM imaging and pseudocoloring. A). Ultrastructure of a biofilm. B). Photoshop overlay to color the different ultrastructures of the biofilm. Scale bar is 20 μm.
Validation of protocol
This protocol has been previously used and validated in the following research article:
Biswas et al. [16]. A Novel Robust Method Mimicking Human Substratum to dissect the Heterogeneity of Candida auris Biofilm Formation. Microbiology Spectrum (Figures 2–4, 6).
General notes and troubleshooting
1. To prevent coverslips from sticking and being damaged while trying to remove them from surfaces, use the lid of a Petri dish and place the coverslips at an angle to prevent them from sticking to the surface completely.
2. It is crucial to use ice-cold water to prevent the formation of gelatin clumps. Clumping can interfere with the coating process and lead to inconsistent results. Therefore, always ensure the water used is sufficiently cold to maintain the gelatin in a smooth, even solution.
3. Carefully following the water washing steps is equally important. Any residual chemicals from previous steps can carry forward to subsequent reactions, potentially preventing proper gelatin coating onto the surfaces. To avoid this, adhere strictly to the washing protocols provided, ensuring thorough and complete rinsing between each step.
4. If you notice that the biofilm remains in a monolayer or has very few cells adhering to the coverslips, this might indicate insufficient washing. In such cases, the number of washing steps with water should be increased. This additional washing can help remove any inhibiting substances and allow for a more robust biofilm formation.
5. It is essential to dry the water completely before proceeding to the next step. Residual moisture can interfere with the reactions, leading to incomplete or failed biofilm formation. Make sure to allow adequate drying time or use appropriate methods to ensure that all water is evaporated before moving forward.
6. Handling the biofilms gently is also critical, especially when pipetting to exchange media or during other manipulations. Biofilms can be fragile, and rough handling can disrupt their structure, leading to inaccurate experimental results. Use gentle pipetting techniques and avoid vigorous movements to maintain the integrity of the biofilms.
7. When using FIJI software for image analysis, avoid maximizing the window. Maximizing the FIJI window can cause the picture to load improperly, which may interfere with image processing and analysis. Keep the window in its default size to ensure smooth operation and prevent software issues.
8. In addition to these specific steps, general troubleshooting tips include ensuring that all equipment and materials are prepared and used correctly. If the coverslips are not fully coated and the biofilm is not forming as expected, revisit each step of the protocol to identify any potential deviations or mistakes. Proper coating and biofilm formation are critical for obtaining reliable results.
9. For imaging issues with both microscopy and SEM, ensure that the biofilm preparation is consistent and the equipment is correctly calibrated. If problems persist, consult the equipment manuals or seek advice from colleagues or technical support.
10. By adhering to these detailed steps and precautions, the quality and reliability of biofilm experiments can be significantly improved. Be meticulous in following protocols and addressing any issues promptly to achieve the best possible outcomes.
Acknowledgments
This work was supported by the Ramalingaswami fellowship grant from the Department of Biotechnology, India and the Science and Engineering Research Board (SERB) DST project funding (EEQ/2022/000606). The protocol described in this paper was published in Microbiology Spectrum (Biswas et al. [16]). A Novel Robust Method Mimicking Human Substratum To Dissect the Heterogeneity of Candida auris Biofilm Formation. 10.1128/spectrum.00892-23.
Competing interests
The authors declare no conflicts of interest.
References
Brouwer, A. E., van Kan, H. J. M., Johnson, E., Rajanuwong, A., Teparrukkul, P., Wuthiekanun, V., Chierakul, W., Day, N. and Harrison, T. S. (2007). Oral versus Intravenous Flucytosine in Patients with Human Immunodeficiency Virus-Associated Cryptococcal Meningitis. Antimicrob Agents Chemother. 51(3): 1038–1042.
Allaw, F., Kara Zahreddine, N., Ibrahim, A., Tannous, J., Taleb, H., Bizri, A. R., Dbaibo, G. and Kanj, S. S. (2021). First Candida auris Outbreak during a COVID-19 Pandemic in a Tertiary-Care Center in Lebanon. Pathogens. 10(2): 157.
Satoh, K., Makimura, K., Hasumi, Y., Nishiyama, Y., Uchida, K. and Yamaguchi, H. (2009). Candida auris sp. nov., a novel ascomycetous yeast isolated from the external ear canal of an inpatient in a Japanese hospital. Microbiol Immunol. 53(1): 41–44.
Garcia-Effron, G., (2020). Rezafungin—Mechanisms of Action, Susceptibility and Resistance: Similarities and Differences with the Other Echinocandins. J Fungi. 6(4): 262.
Abe, M., Katano, H., Nagi, M., Higashi, Y., Sato, Y., Kikuchi, K., Hasegawa, H. and Miyazaki, Y. (2020). Potency of gastrointestinal colonization and virulence of Candida auris in a murine endogenous candidiasis. PLoS One. 15(12): e0243223.
Lee, W. G., Shin, J. H., Uh, Y., Kang, M. G., Kim, S. H., Park, K. H. and Jang, H. C. (2011). First Three Reported Cases of Nosocomial Fungemia Caused by Candida auris. J Clin Microbiol. 49(9): 3139–3142.
Morales-López, S. E., Parra-Giraldo, C. M., Ceballos-Garzón, A., Martínez, H. P., Rodríguez, G. J., Álvarez-Moreno, C. A. and Rodríguez, J. Y. (2017). Invasive Infections with Multidrug-Resistant Yeast Candida auris, Colombia. Emerg Infect Dis. 23(1): 162–164.
Organization, W. H., (2022). WHO fungal priority pathogens list to guide research, development and public health action. World Health Organization.
Cristina, M. L., Spagnolo, A. M., Sartini, M., Carbone, A., Oliva, M., Schinca, E., Boni, S. and Pontali, E. (2023). An Overview on Candida auris in Healthcare Settings. J Fungi. 9(9): 913.
Omardien, S. and Teska, P. (2024). Skin and hard surface disinfection against Candida auris– What we know today. Front Med. 11e1312929.
Cadnum, J. L., Shaikh, A. A., Piedrahita, C. T., Sankar, T., Jencson, A. L., Larkin, E. L., Ghannoum, M. A. and Donskey, C. J. (2017). Effectiveness of Disinfectants Against Candida auris and Other Candida Species. Infect Control Hosp Epidemiol. 38(10): 1240–1243.
De Gaetano, S., Midiri, A., Mancuso, G., Avola, M. G. and Biondo, C. (2024). Candida auris Outbreaks: Current Status and Future Perspectives. Microorganisms. 12(5): 927.
Horton, M. V., Johnson, C. J., Kernien, J. F., Patel, T. D., Lam, B. C., Cheong, J. Z. A., Meudt, J. J., Shanmuganayagam, D., Kalan, L. R. and Nett, J. E. (2020). Candida auris Forms High-Burden Biofilms in Skin Niche Conditions and on Porcine Skin. mSphere. 5(1): e00910–19.
Sherry, L., Ramage, G., Kean, R., Borman, A., Johnson, E. M., Richardson, M. D. and Rautemaa-Richardson, R. (2017). Biofilm-Forming Capability of Highly Virulent, Multidrug-Resistant Candida auris. Emerg Infect Dis. 23(2): 328–331.
Biswas, B., Gangwar, G., Nain, V., Gupta, I., Thakur, A. and Puria, R. (2022). Rapamycin and Torin2 inhibit Candida auris TOR: Insights through growth profiling, docking, and MD simulations. J Biomol Struct Dyn. 41(17): 8445–8461.
Biswas, B., Rana, A., Gupta, N., Gupta, I., Puria, R. and Thakur, A. (2023). A Novel Robust Method Mimicking Human Substratum To Dissect the Heterogeneity of Candida auris Biofilm Formation. Microbiol Spectrum. 11(4): e00892–23.
Article Information
Publication history
Received: Jul 5, 2024
Accepted: Oct 23, 2024
Available online: Nov 20, 2024
Published: Jan 5, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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5,147 | https://bio-protocol.org/en/bpdetail?id=5147&type=0 | # Bio-Protocol Content
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Capacitance Measurements of Exocytosis From AII Amacrine Cells in Retinal Slices
EH Espen Hartveit
MV Margaret L. Veruki
Published: Vol 15, Iss 1, Jan 5, 2025
DOI: 10.21769/BioProtoc.5147 Views: 234
Reviewed by: Wallace B. ThoresonRuth HeidelbergerAkira Karasawa
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Original Research Article:
The authors used this protocol in The Journal of Neuroscience Mar 2022
Abstract
During neuronal synaptic transmission, the exocytotic release of neurotransmitters from synaptic vesicles in the presynaptic neuron evokes a change in conductance for one or more types of ligand-gated ion channels in the postsynaptic neuron. The standard method of investigation uses electrophysiological recordings of the postsynaptic response. However, electrophysiological recordings can directly quantify the presynaptic release of neurotransmitters with high temporal resolution by measuring the membrane capacitance before and after exocytosis, as fusion of the membrane of presynaptic vesicles with the plasma membrane increases the total capacitance. While the standard technique for capacitance measurement assumes that the presynaptic cell is unbranched and can be represented as a simple resistance-capacitance (RC) circuit, neuronal exocytosis typically occurs at a distance from the soma. Even in such cases, however, it can be possible to detect a depolarization-evoked increase in capacitance. Here, we provide a detailed, step-by-step protocol that describes how "Sine + DC" (direct current) capacitance measurements can quantify the exocytotic release of neurotransmitters from AII amacrine cells in rat retinal slices. The AII is an important inhibitory interneuron of the mammalian retina that plays an important role in integrating rod and cone pathway signals. AII amacrines release glycine from their presynaptic dendrites, and capacitance measurements have been important for understanding the release properties of these dendrites. When the goal is to directly quantify the presynaptic release, there is currently no other competing method available. This protocol includes procedures for measuring depolarization-evoked exocytosis, using both standard square-wave pulses, arbitrary stimulus waveforms, and synaptic input.
Key features
• Quantification of exocytosis with the Sine + DC technique for visually targeted AII amacrines in retinal slices, using voltage-clamp and whole-cell patch-clamp recording.
• Because exocytosis occurs away from the somatic recording electrode, the sine wave frequency must be lower than for the standard Sine + DC technique.
• Because AII amacrines are electrically coupled, the sine wave frequency must be sufficiently high to avoid interference from other cells in the electrically coupled network.
• The protocol includes procedures for measuring depolarization-evoked exocytosis using standard square-wave pulses, stimulation with arbitrary and prerecorded stimulus waveforms, and activation of synaptic inputs.
Keywords: AII amacrine cell Capacitance Compartmental model Dendrites Exocytosis Glycine Inhibitory interneuron Patch-clamp recording Presynaptic Retina Sine + DC Slice preparation
Graphical overview
Measuring changes in the membrane capacitance of AII amacrine cells during whole-cell patch-clamp recording in rat retinal slices
Background
In a chemical synapse, a neurotransmitter is released by exocytosis from the presynaptic neuron [1]. For a morphologically discrete synapse, the neurotransmitter diffuses across the synaptic cleft, binds to postsynaptic, ligand-gated ion channels, and typically increases their open probability. This can be measured electrophysiologically as a postsynaptic change in current (voltage clamp) or change in voltage (current clamp). Under ideal conditions, the evoked current will directly represent the underlying conductance change but will only be indirectly related to the magnitude and time course of the presynaptic exocytosis. Because the exocytosis corresponds to the fusion of synaptic vesicles with the presynaptic plasma membrane, the presynaptic capacitance will increase in proportion to the summed capacitance of all released vesicles. The capacitance can be measured with high temporal resolution using a lock-in amplifier, i.e., a phase-sensitive detector, implemented in hardware or software.
Standard capacitance measurement of exocytosis assumes an unbranched cell, represented by a simple RC circuit [2,3]. With the branched morphology of neurons, it is of interest to extend capacitance measurements to such structures [4,5]. Over the last 30 years or so, whole-cell recordings for measuring capacitance have been made directly at different presynaptic boutons where exocytosis takes place, e.g., mossy fiber boutons in the hippocampus [6], goldfish bipolar cell terminals [7], rat rod bipolar cell terminals [8], calyx of Held terminals [9,10], and posterior pituitary gland terminals [11].
Attempts have also been made to measure exocytosis occurring at a distance from the recording pipette, e.g., using somatic recordings of mouse rod bipolar cells with very short axons [12]. More recently, capacitance measurements were extended to AII amacrine cells in mouse retina [13]. The AII is an axonless retinal interneuron with presynaptic dendrites that provide glycinergic synapses onto OFF-cone bipolar cells and OFF-ganglion cells [14]. Using activation of voltage-gated Ca2+ channels to trigger exocytosis, Balakrishnan et al. [13] used capacitance measurements to characterize several important functional properties of the glycinergic synapses. It is a problem for the interpretation of their results, however, that exocytosis in AIIs takes place at a distance from the soma and that these cells are electrically coupled (via gap junctions) to each other and to ON-cone bipolar cells [15,16]. If the goal is to measure the true capacitance increase following exocytosis distributed across several presynaptic dendrites, several constraints apply. First, because exocytosis occurs at a distance from the somatic pipette, the sine wave frequency used to measure the capacitance must be low enough that the electrotonic attenuation from the soma does not exclude some presynaptic terminals from contributing to the measurements. On the other hand, the sine wave frequency must be high enough that attenuation prevents electrotonic transmission through gap junctions that couple to neighboring cells, which can compromise the measurements. Using recently developed compartmental models of AII amacrine cells [17], it was possible to explore these issues computationally and estimate a range of sine wave frequencies that optimizes the trade-off between these conflicting demands [18].
Materials and reagents
Biological materials
1. Rat (Wistar HanTac, Taconic Bioscience)
Reagents
1. Sodium chloride (NaCl) (Sigma-Aldrich, catalog number: 71376 1 kg, CAS number: 7647-14-5)
2. Sodium hydrogen carbonate (NaHCO3) (Sigma-Aldrich, catalog number: S6014 500 g, CAS number: 144-55-8)
3. Potassium chloride (KCl) (Sigma-Aldrich, catalog number: 60128 250 g, CAS number: 7447-40-7)
4. Calcium chloride dihydrate (CaCl2·2H2O) (Sigma-Aldrich, catalog number: 21097 250 g, CAS number: 10035-04-8)
5. Magnesium chloride hexahydrate (MgCl2·6H2O) (Sigma-Aldrich, catalog number: 63064 500 g, CAS number: 7791-18-6)
6. D-Glucose (Sigma-Aldrich, catalog number: G-8280 1 kg, CAS number: 50-99-7)
7. Potassium gluconate (K-gluconate) (Sigma-Aldrich, catalog number G4500 100 g, CAS number: 299-27-4)
8. Potassium hydroxide (KOH) (Sigma-Aldrich, catalog number: 60369 500 g, CAS number: 1310-58-3)
9. Cesium methanesulfonate (CsCH3SO3) (Sigma-Aldrich, catalog number: 368903 25 g, CAS number: 2550-61-0)
10. Cesium chloride (CsCl) (Sigma-Aldrich, catalog number: 1020390050, CAS number: 7647-17-8)
11. Cesium hydroxide (CsOH), 50% (wt) solution in H2O (Sigma-Aldrich, catalog number: 232068 100 g, CAS number: 21351-79-1)
12. Tetraethylammonium chloride (TEA-Cl, (C2H5)4NCl) (Sigma-Aldrich, catalog number: T2265 100 g, CAS number: 56-34-8)
13. 4-(2-Hydroxyethyl)piperazine-1-ethanesulfonic acid (HEPES) (Sigma-Aldrich, catalog number: H3375 100 g, CAS number: 7365-45-9)
14. HEPES, hemisodium salt (hemi-Na salt) (Sigma-Aldrich, catalog number: H7637 100 g, CAS number: 103404-87-1)
15. Ethylene glycol-bis(2-aminoethylether)-N,N,N',N'-tetraacetic acid (EGTA) (Fluka, catalog number: 03778 50 g, CAS number: 67-42-5)
16. Adenosine 5'-triphosphate magnesium salt (MgATP) (magnesium ATP) (Sigma-Aldrich, catalog number: A9187 1 g, CAS number: 74804-12-9)
17. Guanosine 5'-triphosphate sodium salt (Na3GTP) (sodium GTP) (Sigma-Aldrich, catalog number: G8877 100 mg, CAS number: 36051-31-7)
18. Alexa Fluor 594 hydrazide, Na salt (Thermo Fisher Scientific, Invitrogen, catalog number: A10438)
19. Alexa Fluor 488 hydrazide, Na salt (Thermo Fisher Scientific, Invitrogen, catalog number: A10436)
20. (-)-Bicuculline methochloride (HelloBio, catalog number: HB0895 50 mg, CAS number: 38641-83-7)
21. Strychnine hydrochloride (Research Biochemicals Int., catalog number: S-124). For a current source, see Sigma-Aldrich, catalog number: S8753 (25 g, CAS number: 1421-86-9)
22. 6-cyano-7-nitroquinoxaline-2,3-dione disodium salt (CNQX) (HelloBio, catalog number: HB0205 10 mg, CAS number: 479345-85-8)
23. (RS)-3-(2-carboxypiperazin-4-yl)-propyl-1-phosphonic acid (CPP) (HelloBio, catalog number: HB0036 50 mg, CAS number: 100828-16-8)
24. Tetrodotoxin, citrate salt (TTX) (HelloBio, catalog number: HB1035 1 mg, CAS number: 18660-81-6)
25. Ames medium powder [Sigma-Aldrich, catalog number: A1420 (10X1L)]
26. N-(2,6-Dimethylphenylcarbamoylmethyl)triethylammonium chloride (QX314 chloride) (Tocris, catalog number: 2313, CAS number: 5369-03-9)
27. Isoflurane for gas anesthesia (Zoetis Animal Health ApS, catalog number: 002185)
28. Acetone for cleaning platinum-iridium wire before gluing nylon strings onto it to make a slice harp (Merck, catalog number: 1.00014, CAS number: 67-64-1)
29. Sodium hypochlorite (NaOCl) (4% in water, chlorine bleach; can be obtained from the local grocery store)
Solutions
1. Extracellular buffer solution used for dissection (EC3000) (see Recipes)
2. Extracellular bath solution (EC1000) (see Recipes)
3. Intracellular pipette stock solution at 1.25× concentration (IC8503) (see Recipes)
4. Intracellular pipette solution at 1× concentration (IC8503) (see Recipes)
5. Intracellular pipette stock solution at 1.25× concentration (IC4101) (see Recipes)
6. Intracellular pipette solution at 1× concentration (IC4101) (see Recipes)
7. Intracellular pipette stock solution at 1.25× concentration (IC4202) (see Recipes)
8. Intracellular pipette solution at 1× concentration (IC4202) (see Recipes)
9. QX314 (stock solution, 50 mM) (see Recipes)
10. Alexa 594 (stock solution, 1 mM) (see Recipes)
11. Alexa 488 (stock solution, 1 mM) (see Recipes)
12. KCl (stock solution, 1 M) (see Recipes)
13. MgCl2 (stock solution, 1 M) (see Recipes)
14. CaCl2 (stock solution, 1 M) (see Recipes)
15. KOH (to adjust pH, 2 M) (see Recipes)
16. KOH (to adjust pH, 0.2 M) (see Recipes)
17. Ames stock solution (see Recipes)
18. Ames storage (incubation) solution (see Recipes)
19. CNQX (stock solution, 100 mM) (see Recipes)
20. Bicuculline (stock solution, 10 mM) (see Recipes)
21. Strychnine (stock solution, 10 mM) (see Recipes)
22. CPP (stock solution, 50 mM) (see Recipes)
23. TTX (stock solution, 0.3 mM) (see Recipes)
Note: Here and later, the numbers used to identify specific extra- and intracellular solutions are essentially arbitrary and follow a system used in our laboratory (based on the functionality of the Patchmaster software from HEKA Elektronik).
Recipes
1. Extracellular buffer solution used for dissection (EC3000)
Reagent Final concentration Quantity or Volume (for 1 L)
NaCl 145 mM 8.474 g
HEPES (hemi-Na salt) 5 mM 1.247 g
KCl 2.5 mM 2.5 mL of 1 M stock
CaCl2 2.5 mM 2.5 mL of 1 M stock
MgCl2 1 mM 1 mL of 1 M stock
D-Glucose 10 mM 1.802 g
H2O (MilliQ) n/a to 1,000 mL
Total n/a 1,000 mL
Adjust to pH 7.4 with 1 M HCl. Prepare 1,000 mL each time and store at 4 °C. Typically used within a week, keep for up to 10 days.
2. Extracellular bath solution (EC1000)
Reagent Final concentration Quantity or Volume (for 2 L)
NaCl 125 mM 14.610 g
NaHCO3 25 mM 4.2 g
KCl 2.5 mM 5 mL of 1 M stock
CaCl2 2.5 mM 5 mL of 1 M stock
MgCl2 1 mM 2 mL of 1 M stock
D-Glucose 10 mM 3.604 g
H2O (MilliQ) n/a to 2,000 mL
Total n/a 2,000 mL
Prepare 2,000 mL for each experiment. Add all ingredients except CaCl2 to a 2 L volumetric flask. Fill up with H2O but leave enough space for the addition of 5 mL 1 M CaCl2. After all solids have been dissolved and the solution is well mixed, pour into a glass bottle that will be used for the rest of the experiment. Osmolality ~300 mOsm.
Note: Do not add CaCl2 before the solution has been saturated with CO2 (see section F below). If the solution has not been saturated with CO2, Ca2+ will precipitate as CaCO3.
3. Intracellular pipette stock solution at 1.25× concentration (IC8503)
Reagent Final concentration Quantity or Volume (for 1.25× concentration)
CsCH3SO3 80 mM 2.280 g (for 100 mL)
CsCl 40 mM 0.8418 g (for 100 mL)
TEA-Cl 10 mM 0.2071 g (for 100 mL)
HEPES 28 mM 0.8358 g (for 100 mL)
EGTA 2 mM 0.0951 g (for 100 mL)
MgATP 3 mM 0.07134 g (for 40 mL)
Na3GTP 1 mM 0.024654 g (for 40 mL)
CsOH (adjust pH to 7.3) n/a n/a
H2O (MilliQ) n/a n/a
Making a stock solution at 1.25× concentration that gets diluted to a final 1× concentration for the experiment provides flexibility with respect to adding fluorescent dye and specific pharmacological agents. Make up a 40 mL stock solution at 1.25× concentration and store 1 mL aliquots at -20 °C. On the day of the experiment (or shortly before), dilute the 1.25× solution to 1× final concentration by adding water before use. If one decides to also add fluorescent dye (dissolved in water) and/or specific pharmacological compounds (dissolved in water), the volume of water is reduced correspondingly such that the final volume is correct for a 1× solution (see example Recipe below).
In the example Recipe described here, first make up 100 mL of solution at 1.25× concentration containing CsCH3SO3, CsCl, TEA-Cl, HEPES, and EGTA and adjust the pH to 7.3 (with CsOH). From this solution, measure out 40 mL, dissolve the calculated amounts of MgATP and Na3GTP and adjust the pH to 7.3 (only a small amount of CsOH is needed for the second adjustment). Store 1 mL aliquots at -20 °C and dilute to 1× before use.
Caution: CsOH is a very strong base and must be handled with care.
Note: When making up an intracellular pipette solution, there are mutual constraints that influence the accuracy of the different concentrations, the stability of specific compounds, and the total cost of the chemical compounds. On the one hand, preparing a larger volume and adding a larger amount of each chemical increases the accuracy of the concentrations. On the other hand, preparing a smaller volume decreases the total cost. The example here attempts to reach a reasonable compromise and involves preparing a larger initial volume with less expensive compounds, from which a smaller volume is used to prepare the final stock solution. When adjusting the pH (and ideally also the osmolality) of the final solution, there are two challenges. First, volumetric flasks used to prepare solutions with accurate final volumes do not lend themselves to measuring pH using conventional pH electrodes. Second, the base (or acid) that needs to be added cannot be too diluted, as this tends to increase the final volume too much, and also cannot be too concentrated, as it becomes difficult to reach the desired pH without overshooting. One way of handling these problems is to reduce the volume of the solution for which pH is adjusted, approximately by the expected volume (ideally a little less) of base (or acid) whereby pH is adjusted. When the pH has been adjusted to the desired value, the final volume can be checked again in a volumetric flask and, if necessary, H2O can be added.
Note: The water content of ATP and GTP salts varies on a batch-by-batch basis. For consistency, it is therefore recommended to calculate the amounts needed for anhydrous compounds and update the calculations according to the exact water content of a given batch.
4. Intracellular pipette solution at 1× concentration (IC8503)
Reagent Final concentration Quantity or Volume (for 500 μL)
IC8503 at 1.25× 1× 400 μL
Alexa 594 50 μM 25 μL of 1 mM stock
QX314 chloride 2 mM 40 μL of 50 mM stock
H2O (MilliQ) n/a 35 μL
Total n/a 500 μL
After making up 500 μL of intracellular solution at 1× concentration, filter the solution using a 0.22 μm Millex syringe filter. Keep the aliquots on ice during the experiment and freeze at -20 °C between experiments.
Note: Most experimental designs will want to block the Nav channels that mediate spiking in AII amacrine cells for capacitance measurements of exocytosis. One possible solution is to add TTX, a selective blocker of (most types of) Nav channels, to the extracellular bath solution. However, TTX is fairly expensive, and another method is to add the Nav channel blocker QX314, a membrane-impermeable derivative of lidocaine, to the intracellular solution. For a neuron like the AII amacrine cell, Nav channels are blocked within a few minutes after establishing the whole-cell configuration, corresponding to the time it takes for diffusion of QX314 to the subcellular location of the Nav channels.
Note: It is recommended to protect fluorescent dyes from light exposure by covering the corresponding vials with aluminum foil and/or keeping them in a light-tight container.
5. Intracellular pipette stock solution at 1.25× concentration (IC4101)
Reagent Final concentration Quantity or Volume (for 100 mL of 1.25× concentration)
K-gluconate 125 mM 3.6594 g (for 100 mL)
NaCl 8 mM 1 mL of 1 M stock (for 100 mL)
CaCl2 1 mM 0.125 mL of 1 M stock (for 100 mL)
HEPES 10 mM 0.2975 g (for 100 mL)
EGTA 5 mM 0.2378 g (for 100 mL)
MgATP 3 mM 0.05925 g (for 20 mL)
KOH (adjust pH to 7.3) n/a n/a
H2O (MilliQ) n/a n/a
Make up 100 mL of solution with K-gluconate, NaCl, CaCl2, HEPES, and EGTA and adjust pH to 7.3 with KOH. From this solution, measure out 20 mL and add MgATP. Adjust pH to 7.3 with KOH. Store 1 mL aliquots at -20 °C. On the day of the experiment (or shortly before), dilute the 1.25× solution to 1× final concentration by adding water before use. If one decides to also add fluorescent dye (dissolved in water) and/or specific pharmacological compounds (dissolved in water), the volume of water is reduced correspondingly such that the final volume is correct for a 1× solution (see example Recipe below).
Caution: KOH is a very strong base and must be handled with care.
6. Intracellular pipette solution at 1× concentration (IC4101)
Reagent Final concentration Quantity or Volume
IC4101 at 1.25× 1× 400 μL
Alexa 488 100 μM 50 μL of 1 mM stock
QX314 chloride 2 mM 40 μL of 50 mM stock
H2O (MilliQ) n/a 10 μL
Total n/a 500 μL
After making up 500 μL of intracellular solution at 1× concentration, filter the solution using a 0.22 μm Millex syringe filter. Keep the aliquots on ice during the experiment and freeze at -20 °C between experiments.
Note: It is recommended to protect fluorescent dyes from light exposure by covering the corresponding vials with aluminum foil and/or keeping them in a light-tight container.
7. Intracellular pipette stock solution at 1.25× concentration (IC4202)
Reagent Final concentration Quantity or Volume (for 100 mL of 1.25× concentration)
K-gluconate 125 mM 3.6594 g (for 100 mL)
KCl 5 mM 0.625 mL of 1 M stock (for 100 mL)
NaCl 8 mM 1 mL of 1 M stock (for 100 mL)
HEPES 10 mM 0.2975 g (for 100 mL)
EGTA 0.2 mM 0.0951 g (for 100 mL)
MgATP 4 mM 0.1185 g (for 40 mL)
Na3GTP 1 mM 0.0123 g (for 40 mL)
KOH (adjust pH to 7.3) n/a n/a
H2O (MilliQ) n/a n/a
Make up 100 mL solution at 1.25× concentration with K-gluconate, KCl, NaCl, HEPES, and EGTA and adjust pH to 7.3 with KOH. From this solution, measure out 40 mL and add MgATP and Na3GTP. Adjust pH to 7.3 with KOH. Store 1 mL aliquots at -20 °C. On the day of the experiment (or shortly before), dilute the 1.25× solution to 1× final concentration by adding water before use. If one decides to also add fluorescent dye (dissolved in water) and/or specific pharmacological compounds (dissolved in water), the volume of water is reduced correspondingly such that the final volume is correct for a 1× solution (see example Recipe below).
Caution: KOH is a very strong base and must be handled with care.
8. Intracellular pipette solution at 1× concentration (IC4202)
Reagent Final concentration Quantity or Volume
IC4202 at 1.25× 1× 400 μL
Alexa 594 50 μM 25 μL of 1 mM stock
H2O (MilliQ) n/a 75 μL
Total n/a 500 μL
After making up 500 μL of intracellular solution at 1× concentration, filter the solution using a 0.22 μm Millex syringe filter. Keep the aliquots on ice during the experiment and freeze at -20 °C between experiments.
Note: It is recommended to protect fluorescent dyes from light exposure by covering the corresponding vials with aluminum foil and/or keeping them in a light-tight container.
9. QX314 (stock solution, 50 mM)
Reagent Final concentration Quantity or Volume
QX314 chloride 50 mM 10 mg
H2O (MilliQ) n/a 0.67 mL
Total n/a 0.67 mL
MW 298.85 g/mol. Store at -20 °C in 100 μL aliquots.
10. Alexa 594 (stock solution, 1 mM)
Reagent Final concentration Quantity or Volume
Alexa Fluor 594, hydrazide, Na salt 1 mM 1 mg
H2O (MilliQ) n/a 1.32 mL
Total n/a 1.32 mL
MW 758.79 g/mol. Store at -20 °C in 50 μL aliquots.
11. Alexa 488 (stock solution, 1 mM)
Reagent Final concentration Quantity or Volume
Alexa Fluor 488, hydrazide, Na salt 1 mM 1 mg
H2O (MilliQ) n/a 1.75 mL
Total n/a 1.75 mL
MW 570.48 g/mol. Store at -20 °C in 50 μL aliquots.
12. KCl (stock solution, 1 M)
Reagent Final concentration Quantity or Volume
KCl 1 M 7.456 g
H2O (MilliQ) n/a to 100 mL
Total n/a 100 mL
MW 74.55 g/mol. Prepare 100 mL each time, using a 100 mL volumetric flask. Store at room temperature, preferably in the dark. Keep for up to 4 weeks.
13. MgCl2 (stock solution, 1 M)
Reagent Final concentration Quantity or Volume
MgCl2·6H2O 1 M 10.166 g
H2O (MilliQ) n/a to 50 mL
Total n/a 50 mL
MW 203.30 g/mol. Prepare 50 mL each time, using a 50 mL volumetric flask. Store at room temperature, preferably in the dark. Keep for up to 4 weeks.
Note: Please note that MgCl2 is very hygroscopic and will absorb water. Depending on the extent to which this happens, the true amount of salt added will be reduced. To prevent (or minimize) this problem, only purchase relatively small amounts that will be consumed over a reasonable period of time, keep the container tightly closed, and only open the container briefly when weighing out material.
14. CaCl2 (stock solution, 1 M)
Reagent Final concentration Quantity or Volume
CaCl2·2H2O 1 M 14.701 g
H2O (MilliQ) n/a to 100 mL
Total n/a 100 mL
MW 147.01 g/mol. Prepare 100 mL each time, using a 100 mL volumetric flask. Store at room temperature, preferably in the dark. Keep for up to 4 weeks.
Note: Please note that CaCl2 is very hygroscopic and will absorb water. See note for recipe 13 above.
15. KOH (to adjust pH, 2 M)
Reagent Final concentration Quantity or Volume
KOH ~2 M ~100 mg
H2O (MilliQ) n/a 1 mL
Total n/a 1 mL
MW 56.11 g/mol.
Caution: KOH is a very strong base and must be handled with care. Because of potential ion exchange, it is recommended to prepare solutions of KOH in plastic containers (not glassware).
Note: For adjusting pH in intracellular pipette solutions based on K+ salts. KOH comes in the form of pellets, with one pellet weighing approximately 100 mg. To adjust pH, it is useful to have a solution of KOH at approximately 2 M, corresponding to one pellet dissolved in 1 mL of H2O. In addition to the 2 M KOH solution, it is useful to also have a 0.2 M solution of KOH; see recipe below. When adjusting the pH of s small volume of intracellular pipette solution, it is useful to start by adding KOH at a high concentration such that the volume of the solution does not change much. When the pH has almost reached the target value, continuing with the high concentration risks overshooting the target value. Instead, add KOH at the lower concentration (0.2 M).
16. KOH (to adjust pH, 0.2 M)
Reagent Final concentration Quantity or Volume
KOH ~0.2 M ~0.1 mL of 2 M stock solution
H2O (MilliQ) n/a 0.9 mL
Total n/a 1 mL
See comments above for 2 M KOH.
17. Ames stock solution
Reagent Final concentration Quantity or Volume
Ames medium powder n/a 8.8 g (1 glass vial for 1 L)
H2O (MilliQ) to 1,000 mL
Total n/a to 1,000 mL
Prepare 1,000 mL each time and store 50 mL aliquots at -20 °C.
18. Ames storage (incubation) solution
Reagent Final concentration Quantity or Volume
Ames stock solution n/a 50 mL
NaHCO3 25 mM 105 mg (for 50 mL)
Total n/a 50 mL
Thaw a 50 mL aliquot on the day of the experiment. Bubble solution with a gas composed of 95% O2 and 5% CO2 for approximately 20 min (until solution is saturated with CO2). Then, add 105 mg of NaHCO3 and stir until dissolved. Discard the solution after the experiment day.
Note: If NaHCO3 is added before the solution is saturated with CO2, Ca2+ will precipitate as CaCO3.
19. CNQX (stock solution, 100 mM)
Reagent Final concentration Quantity or Volume
CNQX 100 mM 10 mg
H2O (MilliQ) n/a 362 μL
Total n/a 362 μL
MW 276.12 g/mol. Store at -20 °C in 50 μL aliquots.
Caution: CNQX may be toxic and must be handled with care.
20. Bicuculline (stock solution, 10 mM)
Reagent Final concentration Quantity or Volume
Bicuculline methochloride 10 mM 50 mg
H2O (MilliQ) n/a 11.96 mL
Total n/a 11.96 mL
MW 417.85 g/mol. Store at -20 °C in 500 μL aliquots.
Caution: Bicuculline is toxic and must be handled with care.
21. Strychnine (stock solution, 10 mM)
Reagent Final concentration Quantity or Volume
Strychnine hydrochloride × 1.75H2O 10 mM 201.2 mg
H2O (MilliQ) n/a to 50 mL
Total n/a 50 mL
MW 402.38 g/mol (including 1.75 × H2O). Store at -20 °C in 1 mL aliquots.
Caution: Strychnine is toxic and must be handled with care.
22. CPP (stock solution, 50 mM)
Reagent Final concentration Quantity or Volume
CPP 50 mM 50 mg
H2O (MilliQ) n/a 3.96 mL
Total n/a 3.96 mL
MW 252.21 g/mol. Store at -20 °C in 100 μL aliquots.
Caution: CPP may be toxic and must be handled with care.
23. TTX (stock solution, 0.3 mM)
Reagent Final concentration Quantity or Volume
TTX 1 mM 1 mg
H2O (MilliQ) n/a 10.44 mL
Total n/a 10.44 mL
MW 319.27 g/mol. Store at -20 °C in 500 μL aliquots.
Caution: TTX is toxic and must be handled with care.
Laboratory supplies
1. Plastic Petri dish 100 × 15 mm (Corning Inc., catalog number: 351029)
2. Scalpel holder #4 (Fine Science Tools, catalog number: 10004-13)
3. Scalpel blade #20 (Swann Morton Ltd., catalog number: 0086)
4. Scissor, curved, for dissection (B. Braun, catalog number: BC061R)
5. Scissor, small for dissecting eyeball (Fine Science Tools, catalog number: 15000-10)
6. Watchmaker's forceps #5 (VWR, catalog number: 232-1221)
7. Pasteur pipette, with gently fire-polished tip (VWR, catalog number: 612-1709)
8. Borosilicate glass for making patch pipettes (filamented, thick-walled; outer diameter, 1.5 mm; inner diameter, 0.86 mm) (Sutter Instrument, catalog number: BF150-86-10)
9. Parafilm (American National Can, catalog number: 06830)
10. Injection needle, 21 G (Becton, Dickinson and Company, catalog number: 301155)
11. Syringe, 1 mL (Becton, Dickinson and Company, catalog number: 300013)
12. VitraPOR micro-filter-candle tube for bubbling gas in bath solutions, 13 × 25 mm, 8 mm diameter tube, porosity #4 (ROBU Glasfilter-Geraete, catalog number: 18124)
13. Cell strainer, BD Falcon, 100 μm nylon mesh (BD Biosciences, catalog number: 352360)
14. Storage chamber for retinal flatmount pieces (custom-made interface chamber), see section B
15. Plastic box (for making a storage chamber for retinal flatmount pieces, see section B
16. Lens paper (Karl Hecht Assistent, catalog number: 41019010). Cut into small pieces (approximately 15 mm × 5 mm) and store in a small Petri dish
17. Platinum-iridium (Pt-Ir) wire, diameter 0.5 mm, 0.5 mm × 30 cm (World Precision Instruments, catalog number: PTP201)
18. Nylon strings, isolated from nylon stocking
19. Cyanoacrylate (super glue)
20. RTV118 silicone rubber adhesive sealant (Momentive Performance Materials, catalog number: RTV118-85ML)
21. Millex-GV 0.22 μm syringe driven filter tips (Millipore/Merck, catalog number: SLGV004SL)
22. Microloader tips (Eppendorf, catalog number: 5242956.003)
23. Adjustable tubing clamps, "stop-it hose clamp Easy-Click," 10 and 15 mm diameter (Bürkle, catalog number: 8619-0102, 8619-0155)
24. Ag-wire for ground electrodes (patch pipette, bath chamber), Teflon-coated, diameter 0.015" (0.38 mm) (World Precision Instruments, catalog number AGT1510)
25. Small glass beakers, 25 mL (VWR, catalog number: 213-1120)
26. Silicone tubing (thick), ID 5 mm, OD 8 mm (VWR, catalog number: 288-0714)
27. Silicone tubing (thin), ID 2 mm, OD 4 mm (VWR, catalog number: 228-0704P)
28. Tygon tubing (thick), ID 1/16", OD 3/16" (Saint-Gobain Performance Plastics, part number: AAC02002)
29. Tygon tubing (thin), ID 1/16", OD 1/8" (Saint-Gobain Performance Plastics, part number: AAC00002)
Equipment
1. Patch-clamp amplifier (HEKA Elektronik, model: EPC10)
2. Model cell circuit (HEKA Elektronik, model: MC 10)
3. Personal computer for data acquisition and experiment control (Apple Macintosh or Windows PC)
4. Upright, fixed-stage microscope (Olympus/Evident, model BX51WI)
5. Infrared (IR) video camera (TILL Photonics, catalog number: VX55)
6. TV monitor, black/white (CBC Co. Ltd., model CEM-15A)
7. Recording bath chamber insets (aluminum, Teflon-coated) for in vitro slices (Luigs & Neumann, catalog number: 200-100 500 0180-0B), see section C
8. Cover glass (Menzel Gläser), for bottom of recording bath chambers, diameter 50 mm, type #1 (VWR, catalog number: 630-2129), see section C
9. Fluorescence light source for microscope
10. Water immersion objective (×40 or ×60, Olympus/Evident)
11. Dodt gradient contrast (DGC) tube (Luigs & Neumann)
12. Micromanipulator Mini25 motorized (Luigs & Neumann)
13. Fluorescence imaging system (widefield or 2-photon)
14. Vibration isolation table (Technical Manufacturing Corporation [TMC], "Micro-g", model number 63-540)
15. Faraday cage (custom-made)
16. Micro-Osmometer (based on the technique of freezing-point depression to measure osmolality of intracellular pipette solutions) (Fiske Associates, model: 210)
17. Dissection microscope (Leica, model: S6E)
18. Light source for dissection microscope (Volpi, model: Interlux 4100)
19. pH meter (Hanna, catalog number: HI8424)
20. Digital manometer ± 1 psi, incl. custom-made sensor (Sigmann Elektronik, catalog number: 3000703)
21. Water jet pump (BRAND GmbH, catalog number: 1596 00), can be replaced with an electric pump if the use of a water jet pump is not recommended/permitted
22. Pipette puller (Narishige, catalog number: PP-83)
Note: For several items, equivalent commercial alternatives are available. For contrast enhancement, infrared differential interference contrast (IR-DIC) microscopy is an alternative to infrared Dodt gradient contrast (IR-DGC) microscopy.
Software and datasets
1. JPCalcW (Molecular Devices) or JPCalcWin (SDR Scientific), requires license. The Patcher's Power Tools is a free package (required IGOR Pro) that contains some functionality for calculating liquid junction potentials (https://www3.mpibpc.mpg.de/groups/neher/index.php?page=software)
2. Patchmaster v2x92 (HEKA Elektronik/MultiChannel Systems), requires license
3. Fitmaster v2x92 (HEKA Elektronik/MultiChannel Systems), requires license
4. IGOR Pro v9 (WaveMetrics/Sutter Instrument), requires license
Procedure
A. Before experiment day: prepare a U-shaped "harp" to hold retinal slices in perfusion chamber
1. You need Pt-Ir wire, strings isolated from a nylon stocking, and cyanoacrylate super glue (Figure 1).
Figure 1. Custom-made "harp" to hold retinal slices in perfusion chamber. Photo taken during production of a U-shaped slice holder. A flattened piece of Pt-Ir wire is bent into a U-shape and positioned on top of a microscope slide covered with black plastic (for better visibility of the thin nylon strings under the dissection microscope when glued to the Pt-Ir wire). A piece of paper with several black lines spaced 1 mm apart is positioned between the Pt-Ir wire and the black plastic. The black lines serve as a guide when positioning the thin nylon strings isolated from stocking material. The nylon strings are fixed to the Pt-Ir wire using cyanoacrylate glue (visible as the reflective irregular surface on both short arms of the U in the photo). A total of four nylon strings, visible as faint gray lines in the photo, have been stretched across the Pt-Ir wire and fastened with small pieces of tape on either side (left, right). When using the harp to immobilize slices, turn it upside down relative to the orientation in the photo.
2. Cut a piece of wire long enough to be bent into a U-shaped profile with each of the two "arms" approximately 9 mm long and the middle part approximately 11 mm long.
3. After obtaining an adequate shape, use a vice to flatten the wire between two flat pieces of hard metal to change the cross-sectional shape from a circle to approximately a square. Be aware that flattening the wire in this way also slightly increases the length of each part of the U.
4. Clean the U-shaped wire by rinsing it in acetone to remove grease.
5. Place the U on a microscope slide and position it under a dissection microscope.
6. Place single strings isolated from a nylon stocking across both side arms of the U. Fasten the ends of each string with small pieces of tape as you place them. Space the nylon strings approximately 1 mm apart and make sure that they are parallel to each other and the middle part of the U.
7. Use the tip of an injection needle to deposit a small amount of cyanoacrylate glue along the top of each of the side arms of the U to fixate the nylon strings to the Pt-Ir wire. It is best to use very little glue and deposit more than one layer instead of applying too much and risking the glue overflowing and fixing the wire to the microscope slide.
8. When the glue has hardened, the excess nylon strings can be cut at the outside edge of the metal wire using a scalpel blade under the microscope. Start by cutting the strings close to the tape, then turn the metal wire upside-down and cut the strings close to the edge of the metal wire.
9. At the end of each experiment, rinse in distilled water and gently remove small pieces of tissue from the retinal slices using tissue paper.
B. Before experiment day: make an interface storage chamber for retinal tissue
1. You need a clean, empty plastic box made of translucent material (e.g., the plastic box used by Sutter Instrument to store glass capillaries for making patch pipettes), a cell strainer with 100 μm nylon mesh, an injection needle (21 G; Luer fitting), and single-component RTV118 silicone adhesive glue (Figure 2).
Figure 2. Custom-made interface storage chamber for retinal tissue. A. Bottom: Empty storage chamber with insert made from a cell strainer with nylon mesh on top. A. Top: Lid for storage chamber with mounted Luer fit injection needle for gassing solution in chamber. B. Mounted chamber seen from the side. Placing the chamber in a solid plastic base reduces the risk of inadvertent movement of the chamber. C. Mounted chamber filled with Ames storage (incubation) solution seen from the top.
2. Use a scalpel blade to remove the nylon mesh from the sides of the cell strainer, taking care not to damage the mesh on the top.
3. Use RTV118 to glue the bottom of the cell strainer to the bottom of the plastic box. Position the cell strainer a bit to the side to leave room for the gas inlet on the other side.
4. Use a small drilling tool to make a hole for the injection needle in the lid of the plastic box.
5. Use a scalpel blade to cut the injection needle to approximately 15 mm.
6. Insert the injection needle through the hole in the plastic lid and use RTV118 to seal the needle in the lid.
7. Attach a short piece of thin Tygon tubing to the distal end of the injection needle (which will be located inside the box).
8. Verify the volume of aqueous solution needed to fill the chamber such that the nylon mesh at the top of the cell strainer is flush with the fluid level. Small pieces of lens paper with retinal tissue will be positioned at the top of the nylon mesh, with the tissue in contact with the solution below and directly exposed to the atmosphere above (95% O2/5% CO2).
9. Connect the Luer fitting of the injection needle to a source of 95% O2/5% CO2 where the flow can be adjusted.
Note: To prevent growth of microorganisms, sterilize the storage chamber with 70% ethanol after each experiment. Fill the chamber with 70% ethanol, incubate overnight, and rinse the chamber and associated tubing several times with distilled water the next day. Dry before the next experiment.
C. Before experiment day: prepare recording bath chamber
1. Prepare a recording chamber by gluing a round cover glass (50 mm diameter) to the bottom of the chamber with RTV118 silicone rubber adhesive rubber sealant. Take care to apply just the right amount such that little or no silicone mass flows beyond the outer edge of the cover glass and as little as possible silicone mass flows beyond the inner edge. The ideal situation is when no pockets with air are generated at the inner edge of the cover glass, as the bath solution can get trapped and become difficult to rinse and clean after an experiment.
2. Let the silicone harden for 48 h and gently remove any dried silicone rubber that flowed beyond the outer or inner edge of the cover glass. Take care not to break the thin glass of the cover glass. If this happens, remove the glass and silicone rubber and start all over. Take care not to destroy the Teflon coating of the recording chamber; this will compromise the electrical isolation of the solution in the bath chamber and potentially lead to ground loops and increased electrical noise during the electrophysiological recording.
3. A recording chamber prepared in this way can last for several years before it needs to be replaced (e.g., if the cover glass breaks).
D. Before experiment day: chloride Ag-wires for ground electrodes
1. Cut adequate lengths of Ag-wire to be used as ground electrodes for the patch pipette and bath chamber.
2. Remove the Teflon coating from the piece of Ag-wire. After this step, handle the Ag-wire only with clean forceps to avoid contaminating the metal surface with grease from your fingertips. If this still happens, clean the wire in acetone before continuing. Acetone evaporates quickly but make sure that this is the case before continuing to the next step.
3. Connect the Ag-wire to the patch pipette holder or ground electrode holder, following the manufacturer's instructions. Some designs require soldering, while others fasten the wire mechanically without soldering.
4. Immerse the distal (furthest from the holder) length of the Ag-wire in sodium hypochlorite (chlorine bleach) for ≥24 h. This generates a layer of AgCl on the immersed surface of the wire. After successful chloriding, the wire will display an even, light gray coating.
Note: Chlorine bleach is corrosive and must be handled with care.
E. Before experiment day: prepare intracellular solution at 1.25× concentration
1. Prepare intracellular pipette solution (IC8503) at 1.25× concentration (see Recipes).
F. Experiment day: prepare extracellular solution
1. Prepare extracellular bath solution EC1000.
a. Prepare 2 L of EC1000 (see Recipes) and place a glass bottle with the solution at the recording setup.
b. Position a micro-filter-candle tube ("gas bubbler") in the EC1000 solution and start the flow of 95% O2/5% CO2. Connect the gas bubbler to the source of 95% O2/5% CO2 with thick silicone tubing and use a large adjustable clamp to regulate the flow.
c. After bubbling the EC1000 for approximately 30 min (to lower the pH), add 5 mL of 1 M CaCl2.
2. Divide the prepared volume of EC1000 in two parts: one larger (e.g., 1.8 L) to be used as is without any added pharmacological agents, and one smaller (e.g., 0.2 L) to which pharmacological agents are added to block synaptic transmission. Keep the two solution volumes in separate (clearly labeled) bottles and make sure that each is bubbled adequately with 95% O2/5% CO2.
a. To the smaller volume (0.2 L), add the following pharmacological agents:
10 μM CNQX (20 μL of 100 mM stock solution)
10 μM bicuculline (200 μL of 10 mM stock solution)
1 μM strychnine (20 μL of 10 mM stock solution)
20 μM CPP (80 μL of 50 mM stock solution)
Note: To block Na+ channels in AII amacrine cells, one can either add QX314 to the intracellular pipette solution or TTX to the extracellular bath solution. Adding QX314 intracellularly does not block Nav channels in other cells in the tissue, including other AIIs electrically coupled to the cell from which the recording is made. Adding TTX extracellularly blocks all relevant Nav channels but is more expensive than using QX314. To add TTX at a final concentration of 1 μM of the bath solution specified above, add 333 μL of 0.3 mM stock solution for each 100 mL of bath solution.
Note: Several of the pharmacological agents listed above are strong toxins and must be handled with caution.
G. Experiment day: prepare intracellular solution (1×) from 1.25× stock
1. Thaw an aliquot of frozen (-20 °C) solution (IC8503) at 1× concentration. Alternatively, prepare a new solution at 1× concentration from 1.25× stock solution if needed (see Recipes).
2. Keep the intracellular solution on ice for the duration of the experiment. Store frozen at -20 °C between experiments.
H. Prepare Ames storage (incubation) solution for storage of retina tissue ex vivo
1. Thaw an aliquot (50 mL) of frozen (-20 °C) Ames stock solution.
a. Pour the thawed solution into the storage chamber.
b. Bubble solution with gas mixture containing 95% O2/5% CO2.
c. Weigh 105 mg of NaHCO3.
d. After the Ames stock solution has been saturated with 5% CO2 (approximately 20 min) and pH has stabilized, add the NaCO3 to the solution. Make sure that NaHCO3 has been fully dissolved and that the buffered Ames storage (incubation) solution is well mixed.
e. Keep the storage chamber closed.
I. Pull patch pipettes
1. Before the experiment day, lightly fire polish (in a gas flame) both ends of the pipette blanks from which the patch pipettes will be pulled (this protects plastic parts, O-rings, and rubber gaskets of the pipette holder).
2. On the experiment day, use a two-stage pipette puller to pull pipettes for whole-cell recording. When filled with intracellular pipette solution and positioned in the extracellular bath solution, the resistance should be 5–11 MΩ. With the Narishige PP-83 puller, we typically use values for NO.1 and NO.2 HEATER ADJ. of 13.5 and 8.3, respectively. The settings will naturally differ for other pipette pullers.
3. Keep the pipettes in a dust-free container until use. Use pipettes on the same day they were pulled to minimize the accumulation of small dust particles at the pipette tips during storage.
J. Prepare dissection area and tools
1. Prepare and have the following equipment and supplies ready:
a. Small pieces of lens paper.
b. Scalpel blade mounted on scalpel holder.
c. Small curved scissor (for dissection of eyeball from orbit).
d. Iris scissor (for opening the eyeball).
e. Watchmakers forceps × 2.
f. Injection needle mounted on a 1 mL syringe.
g. Small beaker (for rinsing eyeball).
h. Petri dish (100 × 15 mm) filled with cold extracellular buffer solution for dissection (EC3000).
i. Pasteur pipette with gently fire-polished tip.
K. Isolate retina tissue from experimental animal
1. Prepare a Petri dish for dissection.
a. Place the Petri dish under the dissection microscope.
b. Fill the Petri dish with cold EC3000 (let EC3000 stored at 4 °C sit out at room temperature for approximately 1 h before use).
2. Anesthetize and kill rat.
Note: All procedures must be approved and authorized by the local animal welfare authority.
a. Place the rat in the chamber for gas anesthesia.
b. Start the flow of 100% O2 through the anesthesia chamber.
c. After approximately 10 min, add isoflurane at a concentration of 2%–4%.
d. When the rat is unconscious, check reflexes for depth of anesthesia.
e. When anesthesia is sufficiently deep, remove the rat from the chamber and perform cervical dislocation to kill it.
3. Dissect out both eyes.
a. Quickly dissect out both eyes using a curved scissor.
b. Rinse eyes in a small beaker with cold EC3000.
c. After rinsing, place the eyeball in a Petri dish filled with EC3000 under the dissection microscope.
4. Isolate the retina by dissecting it from the eyeball (this must be performed for both eyes in rapid succession; alternatively, two people need to collaborate at this stage, with each person dissecting one eye). The procedure for dissecting the eye, as well as cutting slices, is described stepwise below and schematically illustrated in Figure 3A.
Figure 3. Dissecting the eye, storing retinal tissue, and perfusion and visualization of retinal slices. A. Procedure for dissection of the eye, starting with an encircling cut through the three layers of the eyeball (sclera, choroid, and retina) just behind the limbus (border between the cornea and sclera). After removing the cornea and lens, the remaining eye cup contains sclera, choroid, and retina. The retina is then isolated as a "retina cup" by detaching it from the posterior eye cup. The retina is cut into four quadrants (kept in the storage chamber) and, subsequently, each quadrant is cut into a number of vertical slices after trimming the quadrant into the shape of a rectangle. B. Pieces of retinal tissue ("quadrants") in the storage chamber, each isolated piece is positioned on a small piece of lens paper that sits on top of the nylon mesh of a cell strainer. The storage chamber is designed such that the nylon mesh is located at the interface between the Ames storage (incubation) solution and the atmosphere (95% O2/5% CO2) above. C. Schematic view of the recording chamber seen from above. The retinal slices are positioned on their side on the glass bottom of the chamber and stabilized by a "harp" made of a U-shaped, flattened Pt-Ir wire with parallel thin nylon strings attached to the frame with cyanoacrylate glue. Note the position in the recording chamber of the inflow of extracellular bath solution and the removal of solution at the opposite end (outflow). The position of the reference/ground electrode (Ag-AgCl wire) should be close to the outflow to avoid contamination of the bath solution. D. Cut surface of the retinal slice in the recording chamber positioned on the microscope stage and visualized with infrared Dodt gradient contrast. The retinal layers can be clearly visualized (photoreceptor layer at top, ganglion cell and nerve fiber layer at bottom) and are indicated by abbreviations (PhR, photoreceptors; ONL, outer nuclear layer; OPL, outer plexiform layer; INL, inner nuclear layer; IPL, inner plexiform layer; GCL, ganglion cell layer; NFL, nerve fiber layer). The cell bodies of AII amacrine cells are located at the border between the INL and the IPL.
a. Place the Petri dish with the eyeball under the dissection microscope and locate the eyeball in the oculars.
b. Grasp extraocular tissue with watchmaker's forceps and use a small scissor (iris scissor) to remove larger pieces of extraocular tissue (connective tissue, fat, and extraocular muscles).
c. When the eyeball has been cleaned, use a sharp injection needle mounted on a 1 mL syringe to pierce the wall of the eyeball, approximately at the equator.
d. Using a watchmaker's forceps to stabilize the eyeball, insert one prong of the small iris scissor in the small hole of the eyeball and make a continuous cut along the equator of the eyeball to separate the front [cornea plus anterior half of the sclera from the posterior half of the sclera (the cut shall be approximately along the location of the ora serrata)].
e. Remove the front half (cornea and sclera, including the lens). The remaining posterior part will be referred to as the eye cup.
f. Using one watchmaker's forceps to stabilize the eye cup, use the other watchmaker's forceps to gently remove the vitreous from the eye cup. Be careful not to touch the retina with the tip of the forceps. Instead, close the forceps as closely as possible to the retina and pull the forceps away. Continue until no more vitreous can be removed.
g. Gently remove the retina from the remaining choroid and sclera. One technique involves inserting a blunt probe (made by melting the tip of a Pasteur pipette) below the retina and moving gently sideways. An alternative technique involves using two watchmaker's forceps to grab small (peripheral) regions of the sclera and pull them apart to detach the retina. When most of the retina has been detached from the choroid, use the small iris scissor to cut the optic nerve as it passes through the lamina cribrosa at the back of the eye. The retina is now isolated as a retina cup.
h. Using a scalpel with a curved blade, divide the retina cup into four approximately equal quadrants. First, place a single cut through the optic disk of the retina cup. Then divide each half in two by placing a second cut, also passing approximately through the location of the optic disk and orthogonal to the first cut.
i. With each retina quadrant oriented with ganglion cell side up (photoreceptor side down), use a watchmaker's forceps to gently position a small piece of lens paper under each retina quadrant, lift it out of the solution in the Petri dish, and position the lens paper on the nylon mesh on top of the cell strainer in the storage chamber. The lens paper (with retina quadrant) should be located at the interface between the Ames storage (incubation) solution and the atmosphere inside the storage chamber (Figure 3B).
j. Repeat the procedure above such that all four quadrants (eight when both eyes have been dissected) are located in the storage chamber. Close the lid of the storage chamber.
k. Check that the flow of 95% O2/5% CO2 in the storage chamber is adequate.
l. Let the retina tissue rest for a minimum of 1 h (at room temperature) before starting the preparation of slices and electrophysiological recording.
L. Cut retinal slices
1. Transfer a retina quadrant from the storage chamber to the Petri dish filled with EC3000 (at room temperature). If there is a concern about the state of the tissue or the subsequent procedure takes a long time, the EC3000 solution can be bubbled with 100% O2. If one prefers to prepare the slices in a solution with pH buffered by bicarbonate-CO2 (instead of HEPES), fill the Petri dish with EC1000 (instead of EC3000) and bubble with 95% O2/5% CO2.
a. During the transfer, make sure that the lens paper with retina tissue enters the solution in the Petri dish upside down, i.e., with the retina facing down. If not, the piece of retina often curls up with a small bubble of air "trapped" in the tissue, which can be hard to remove without damaging the retina.
b. Gently remove the piece of lens paper (use a pair of watchmaker's forceps).
2. Cut slices (see schematic in Figure 3A).
a. Trim the retinal quadrant to a strip of tissue of approximately 2 × 5 mm.
b. Grasp one corner of the strip of retinal tissue with the tip of a watchmaker’s forceps and cut vertical slices (parallel to the long axis of the photoreceptors) by hand using a scalpel with a curved blade. Cut the slices in parallel with the short edge (opposite from the corner held by the forceps). The retina is approximately 200 μm thick, meaning that a successfully cut slice will be 100–200 μm thick. Cut 10–15 slices from each quadrant.
3. Transfer the retinal slices to the recording chamber.
a. Using a fire-polished Pasteur pipette, gently suck up one or more of the retinal slices from the Petri dish and gently expel them into a recording chamber (Figure 3C).
b. With all slices in the recording chamber, use two watchmaker's forceps to arrange the slices parallel with each other, making sure all have the same orientation. When a slice is cut at adequate thickness, it usually will lie naturally on its cut side. In this position, a slight curvature reveals which edge corresponds to the photoreceptor side ("outside curve") and which corresponds to the ganglion cell side ("inside curve"). Adequate orientation of the retinal slices in the recording chamber is important for subsequently searching the slices for AII amacrine cells during the recording phase (see below).
c. When the slices are adequately aligned in the recording chamber, gently position the Pt-Ir harp on top of the slices to secure them in place. The parallel nylon strings (glued to the Pt-Ir wire) shall be oriented orthogonally to the long axes of the slices. When the spacing of the nylon strings on the harp and the length of the slices are adequate, each slice will be covered by two or three nylon strings (Figure 3C).
d. Place the recording chamber with retinal slices under the microscope. Make sure that the chamber is well fastened and stabilized.
Note: The slices should be oriented such that when they are viewed on a TV monitor or computer monitor, they appear as in Figure 3D.
Note: A single batch of slices should not be used for more than 3–4 h before being replaced by a new batch.
M. Start perfusion of the recording chamber with retinal slices
1. Start perfusion of the recording chamber.
a. Put the upstream end of the inlet tubing (thick Tygon) in the reservoir with extracellular bath solution (EC1000) saturated with 95% O2/5% CO2. The reservoir with bath solution can conveniently be placed on top or inside (e.g., on a shelf) of a Faraday cage (surrounding the setup to shield the patch-clamp preamplifier/headstage from electrical noise). Fill the inlet tubing going from the reservoir to the recording chamber with EC1000, position the downstream end of the inlet tubing (thin Tygon) into the recording chamber, and start the flow. Start the pump (water suction or electric) and position the upstream end of the outlet tubing (thin silicone) into the recording chamber.
b. With a drop chamber inserted along the course of the inlet tubing (between thick and thin Tygon), adjust the flow to an adequate rate (e.g., 1.5–3 mL/min). To adjust the flow rate, we use an adjustable clamp attached to the thin Tygon tubing a short distance downstream of the drop chamber ("stop-it hose clamp Easy-Click," see Laboratory supplies).
Caution: Too-low flow can compromise oxygen levels and lead to a basic pH in the chamber due to the loss of CO2 to the atmosphere. This can compromise normal cellular physiology and can cause precipitation of Ca2+ (as CaCO3) in the recording chamber and on the microscope objective. Too-high flow can compromise mechanical stability.
2. Position the ground electrode into the recording chamber.
a. Preferentially position the ground electrode such that the location of the holder does not interfere with the positioning and removal of a recording patch pipette.
b. Preferentially position the ground electrode such that it is closer to the suction tip of the outlet tubing through which solution is removed from the recording chamber.
N. Find and record from an AII amacrine cell
1. Prepare the microscope.
a. Mount the water immersion objective on the microscope.
b. Lower the objective into the solution in the recording chamber. Make sure that no air bubbles are trapped below the tip of the objective.
c. Turn on the illumination for the IR-DGC system. Focus on locating the slices on the TV/computer monitor. Once the top surface of the slices has been located, move the focus to one end of the top slice.
d. Use the micromanipulator for the microscope stage/bath chamber holder (manual or motorized) to move the preparation and search systematically along the long axis of each retinal slice for the cell body of an AII amacrine cell.
2. Searching for an AII amacrine cell.
a. The cell bodies of AII amacrine cells are located in the inner nuclear layer toward the border between the inner nuclear layer and the inner plexiform layer, with a thick apical dendrite emanating from the cell body and descending into the inner plexiform layer (Figure 4A, B).
b. When a putative AII amacrine cell has been located, raise the objective approximately 1.5 mm above the position where the focus was on the top surface of the retinal slice. Be careful not to break the contact between the bottom of the objective and the bath solution.
Figure 4. Targeting AII amacrine cells in the retinal slice preparation. A. Video micrograph of retinal slice visualized with IR-DGC microscopy. Note the cell body and thick apical dendrite of an AII amacrine cell (indicated by arrow), with the cell body in the inner nuclear layer (top) and the apical dendrite descending into the inner plexiform layer (bottom). Retinal layers are indicated by abbreviations (INL, inner nuclear layer; IPL, inner plexiform layer; GCL, ganglion cell layer). Scale bar (A, B): 10 μm. B. Same as in A, after establishing a whole-cell recording with a patch pipette positioned on the cell body of the AII (right). C. AII amacrine filled with Alexa 594 during whole-cell recording (different from A, B). Maximum intensity projection generated from widefield fluorescence image stack after deconvolution. Scale bar: 10 μm. D. Characteristic action currents (escape from voltage clamp) evoked in an AII amacrine during whole-cell recording in response to 5 ms depolarizing voltage pulses from Vhold = -60 to -55 mV (voltage stimulus at top). Traces show responses recorded immediately after breaking into the cell (middle) and approximately 3 min later, after diffusion of QX314 from the intracellular pipette solution and block of voltage-gated Na+ (Nav) channels (bottom; n = 4 responses in each condition).
3. Coat the patch pipette with Parafilm and fill it with intracellular solution.
a. Cut a thin strip (e.g., 1 × 30 mm) of Parafilm and wrap it around the tip of the patch pipette. Start as close as possible to the distal tip of the pipette and wrap and stretch the Parafilm around the glass while moving away from the tip toward the "shoulder" and unpulled part of the pipette glass. Make sure that the length of the pipette that will be immersed in the fluid of the bath chamber is covered by Parafilm. Press firmly at the last end of the Parafilm to keep it from unraveling.
b. Fill the tip of the patch pipette with a few microliters of intracellular solution using a syringe with a long, thin tip made by melting the tip of a 1 mL syringe in a gas flame and drawing out the melted tip to a thin tube (alternatively, use a commercial Microloader tip).
c. Mount the patch pipette in the pipette holder (attached to the micromanipulator).
d. Apply positive pressure to the pipette (typically 5–15 mbar) through the suction tubing (thin silicone) attached to the side port of the patch pipette holder and close the valve to maintain the pressure.
4. Move the pipette tip toward the targeted cell.
a. Move the tip of the pipette into the bath solution (with the objective raised to approximately 1.5 mm above the surface of the slice).
b. Move the tip of the pipette under the objective and try to locate it under the microscope. Search for the pipette tip by moving only the pipette. Move the pipette far enough so that the tip will have crossed the vertical midline orthogonal to the horizon of the field of view. Once this has been achieved, only move the pipette sideways, not up and down (as this risks moving the pipette into the slices or the bottom of the bath chamber). Do not move the bath chamber. Unless you are lucky and the pipette tip is in focus, you will observe the pipette movement as a shadow moving across the field of view. When this happens, make small movements with the pipette micromanipulator to figure out if you need to move the pipette out or in, and if you need to move the objective up or down, to bring the tip of the pipette into focus.
c. Once the pipette tip is in focus, lower it to a focal plane just above the top surface of the slice (to avoid breaking the tip of the pipette or moving it accidentally into the slice). Do this by first moving the objective down a small distance, then moving the pipette into focus, and then moving the objective down again, etc.
5. Set up and adjust the test pulse, liquid junction potential, and offset.
Note: The specific details of this protocol are based on the use of the EPC10 patch-clamp amplifier and the Patchmaster software, using the built-in LockIn extension that adds the functionality of a lock-in amplifier implemented in software. This extension must be activated in the Patchmaster Configuration window before using it the first time. Most, but not all functions will have equivalents in corresponding hardware and software from other vendors.
a. Enter the calculated (or measured) value of the liquid junction potential in the Patchmaster software. Patchmaster expects the convention where the liquid junction potential is the potential of the bath with respect to the potential of the pipette [19,20]. The convention is essentially arbitrary, and the particular choice might seem odd, but simplifies subsequent calculations (performed automatically by the Patchmaster software). For the particular combination of extracellular bath solution (EC1000) and intracellular pipette solution (IC8503) used here, the liquid junction potential has been calculated to be +7.3 mV.
b. Once the pipette tip is in the solution of the bath chamber, start the test pulse (e.g., 5 mV amplitude, 5 ms duration) in the software. Click the button SETUP in the Patchmaster amplifier window.
c. Adjust the voltage offset to zero the baseline current.
d. Read off and store the pipette resistance in the software (for documentation purposes).
6. Establish a GΩ-seal and the whole-cell recording configuration.
a. Under continuous visual observation on the TV monitor, position the tip of the patch electrode gently on top of the cell body of the targeted AII amacrine cell. With adequate contrast enhancement (IR-DGC or IRDIC), this can be seen as a small depression on the top of the cell body ("dimpling").
b. Release the positive pressure and apply gentle suction (by mouth).
c. Monitor the resistance and wait for the establishment of a GΩ-seal. If necessary, apply gentle suction and/or hyperpolarization of the patch pipette potential to facilitate the formation of a seal.
d. When a GΩ-seal (≥2 GΩ) has been obtained, neutralize the pipette capacitance using the C-fast circuitry of the amplifier.
e. Set the holding potential to the desired potential, e.g., -60 or -65 mV.
f. Establish the whole-cell configuration by applying suction in combination with brief (e.g., 0.1 ms), high-amplitude (e.g., 400 mV) voltage pulses (actuated by pressing the ZAP button in the Patchmaster amplifier window). Obtaining the whole-cell recording can be observed by the appearance of large-amplitude capacitive charging transients at the onset and offset of the test pulse, as well as by an increase of the steady-state current amplitude during the constant phase of the voltage test pulse.
g. Neutralize the cell's (apparent) capacitance and estimate the series resistance by using the C-slow circuitry of the amplifier. If required, fine-tune the automatic adjustments manually.
h. To minimize the effective capacitance of the pipette, keep fluid levels in the bath chamber as low as possible.
7. Test for "electrophysiological signature" of AII amacrine cells.
a. From a voltage-clamp holding potential of -60 or -65 mV, apply a brief depolarization of 5 to 10 mV. If the recording is indeed from an AII amacrine cell, the depolarization evokes an action current, corresponding to an unclamped action potential [21] at the so-called axon initial segment-like process of the AII (Figure 4D). The final verification must by necessity be morphological (Figure 4C), but it is not recommended to use ordinary widefield fluorescence microscopy until the end of the recording, as even brief exposures can cause phototoxicity. If the experiment is performed in combination with 2-photon microscopy, the risk of phototoxicity is much lower and the cell can safely be visualized immediately to confirm its identity.
8. Monitor pipette pressure in the whole-cell configuration.
a. It has been reported that a small increase in hydrostatic pressure can (reversibly) inhibit compensatory endocytosis [22,23]. It can therefore be an advantage to maintain a slight negative pressure on the pipette after establishing the whole-cell configuration (e.g., -0.7 mbar).
9. Apply the stimulus protocol and acquire data, using the graphical user interface of Patchmaster.
O. Designing a stimulation protocol for an experiment with capacitance measurement of exocytosis
This should be done before the day of the experiment, but for convenience is described here. Designing an optimal voltage-clamp stimulus can be complicated and should ideally be done by testing the patch-clamp amplifier with a model cell that mimics the electrical circuits corresponding to "pipette in bath," "GΩ-seal," and "whole-cell configuration."
Critical: When designing stimulus and acquisition protocols for the amplifier hardware/software, it is crucial to thoroughly test and verify the performance using an electronic model cell (supplied by the amplifier manufacturer or custom-built) before performing an experiment using real cells.
1. To increase the accuracy of capacitance measurements of exocytosis, make sure to manually calibrate the phase shift and attenuation caused by the instrumentation (see section S).
2. When designing the voltage clamp stimulus ("sequence"), take the following points into consideration:
a. The sine wave frequency can in principle range from 100 Hz to 10 kHz. In the examples illustrated below, the frequency is set to 2 kHz, but an analysis using compartmental modeling of AIIs indicates that this frequency clearly underestimates the total increase in capacitance when exocytosis also takes place at more distal lobular dendrites (for details, see [18]).
b. On the one hand, the sine wave amplitude should be as large as possible to increase the signal-to-noise ratio of the measurements.
c. On the other hand, the sine wave amplitude must be small enough that it does not activate voltage-gated currents, i.e., during the application of the sine wave stimulus only the passive leak current should contribute to the evoked current.
d. In addition to the sine wave amplitude, the average potential during the application of the sine wave stimulus will determine whether or not voltage-gated currents are likely to be activated. The average potential should not be so hyperpolarized that the stability of the cell is compromised and not so depolarized that it activates voltage-gated currents. For a sine wave stimulus, one must take these points into consideration when selecting both the sine wave amplitude and the average potential from which it is applied.
e. To evoke exocytosis, the voltage stimulus must contain a depolarizing voltage pulse with an amplitude and duration that is sufficient to activate voltage-gated Ca2+ channels (Cav channels).
f. The cutoff frequency of the lowpass filter applied to the current signal should be set to 2 × fsine, where fsine is the frequency of the sine wave stimulus.
g. The sampling frequency (the inverse of the sampling interval) of the current signal should be set to 10 × fsine.
h. The lock-in calculations of the Patchmaster software provide measurements of Cm (membrane capacitance), Gm [membrane conductance; = 1/Rm (inverse of membrane resistance)], and Gs [series conductance; = 1/Rs (inverse of series resistance)].
i. The default mode of operation of the software implementation of a lock-in amplifier by the EPC10 + Patchmaster instrumentation calculates one data point per sine wave cycle.
3. Example parameters of a voltage stimulus [sequence configured in the graphical editor for a given "Pulse Generator File" (PGF), henceforth referred to as a PGF sequence or PGF for short].
a. To measure depolarization-evoked changes of Cm, Rm, and Rs (ΔCm, ΔRm, ΔRs), presumably reflecting Ca2+-dependent exocytosis, calculate the baseline as the average during a 400 ms period before the stimulus and the response as the average during a 400 ms period after the stimulus (for stimulus durations < 400 ms). For stimulus durations > 400 ms, calculate the baseline as the average during a 1,000 ms period after the stimulus.
b. To activate Cav channels, apply a depolarization from Vhold to -10 or -20 mV. To avoid rundown and sequence effects, do not apply the stimulus more frequently than every 30 s (unless the goal is to study, e.g., vesicle depletion).
c. Configure the voltage stimulus as follows in the PGF editor of Patchmaster (Figure 5):
Sampling interval: 50 μs (20 kHz)
Recording mode: Voltage Clamp
DA: send to Stim1 (send stimulus to Amplifier 1), apply StimScale, use for LockIn
AD: Imon-1 (A), Compression 1 (single sample, 2-byte integer)
LockIn_CM (F), Compression 10 (single sample, 4-byte real)
LockIn_GM (S), Compression 10 (single sample, 4-byte real)
LockIn_GS (S), Compression 10 (single sample, 4-byte real)
FilterFactor: set to a value that results in a lowpass filter cutoff frequency two times larger than fsine
Number of segments: 5
Segment 1: sine wave stimulus, Vhold, 2,000 ms
Segment 2: constant, Vhold, 20 ms
Segment 3: constant, depolarization to -10 or -20 mV (e.g., 100 ms)
Segment 4: constant, Vhold, 100 ms // to let the membrane conductance return to baseline
Segment 5: sine wave stimulus, Vhold, 10,000 ms
Note: It is necessary to add a segment with constant voltage (segment 4 in Figure 5) immediately after a stimulus designed to trigger Ca2+-dependent exocytosis (segment 3 in Figure 5) to allow the evoked change in conductance to return to the resting/baseline conductance before application of a new sine wave voltage stimulus to measure the post-stimulus capacitance.
Figure 5. Voltage stimulus used to measure capacitance before and after a depolarizing pulse. Top: Voltage stimulus designed as described in step O3c with five segments (1–5), including a sine wave stimulus to measure baseline capacitance (1; 2 kHz, ±20 mV from Vhold = -90 mV, 2,000 ms), a constant segment (2; Vhold, 20 ms), a depolarizing pulse to activate voltage-gated Ca2+ channels (3; -20 mV, 100 ms), a constant segment to let the membrane conductance return to baseline (4; Vhold, 100 ms), and a sine wave stimulus to measure capacitance after exocytosis (5; 2 kHz, ±20 mV from Vhold, 12,000 ms). For clarity, only subsegments before and after the depolarizing pulse are displayed. Note that because the sine wave frequency is 2 kHz, individual cycles cannot be resolved at this time scale. Bottom: As in the top graph, but for illustration purposes, the sine wave frequency has been reduced to 5 Hz.
d. Decide the value for Vhold.
During the recording, an AII amacrine cell can be held at a holding potential of -65 mV. However, during the application of the sine wave stimulus, Vhold must be at a more negative potential. It has been reported that Cav channels in (mouse) AII amacrines activate at approximately -55 mV [24]. For a sine wave stimulus of ±15 mV, this means that Vhold must be -75 mV or more negative. For a sine wave stimulus of ±30 mV, this means that Vhold must be -90 mV or more negative. We typically use Vhold = -90 mV and a sine wave stimulus of ±15 mV or ±20 mV.
e. Configure the sine wave as follows in the Sinewave Parameters window of the Patchmaster software. To bring up this window, click the button labeled Sine Wave in the Pulse Generator File window of Patchmaster.
Note: Here and later, text following "//" is commentary.
"Use as LockIn SineWave"
Peak ampl. [mV]: 20 (value) // "value" indicates that the numerical value can be modified
// online "Peak amplitude" is amplitude from baseline, the total // amplitude (peak-to-peak) will be twice as large
Requested frequency: 2.0 kHz
Actual freq.: 2.0 kHz // may differ from "Requested freq."
Points / Cycle: 10
Cycles to Skip: 1 // setting to "1" discards the data points of the first cycle to avoid "swing
// in" effects
Cycles to Average: 1 // setting to "1" provides one measurement point for each cycle
Total Cycles: 24440
V-reversal (mV): -15 mV // the estimated value of the reversal potential of the leak current,
// the exact value is not very critical for the estimation of Cm, Rm,
// and Rs
Note: Configure a voltage stimulus (PGF sequence) to enable leak subtraction. Leak subtraction involves the application of a scaled-down version of the pulse protocol in a voltage range where voltage-gated channels are not active. The resulting current is averaged, scaled, and subtracted from that evoked by the main pulse protocol. This will ideally remove both linear leak currents and capacitive currents [25]. If there is no need to visualize or analyze the voltage-gated Ca2+ current evoked by the depolarizing pulse (corresponding to Segment 3 in the example above), there is no need to add leak subtraction stimuli to the sine wave voltage stimulus. If the user decides to do leak subtraction, however, a standard implementation can be a challenge for an efficient execution of the experiment. The total duration of the sine wave voltage stimulus described earlier is more than 12 s and, with such a long duration, applying an adequate number of leak pulses (larger than or equal to the amplitude of the main pulse divided by the amplitude of the leak pulse) for each ordinary ("non-leak") sweep becomes very time-consuming. However, if the primary goal of the leak subtraction is to visualize and analyze the Cav current, there is no need to apply a leak pulse stimulus that has the same duration as the full stimulus waveform. Instead, it is better to generate a stimulus that only encompasses the three middle segments (2, 3, and 4), i.e., the depolarizing voltage pulse (appropriately scaled) with the two flanking segments.
4. Example parameters of a voltage stimulus (PGF sequence) to estimate the leak conductance.
a. Configure the voltage stimulus as follows in the PGF editor of Patchmaster (Figure 6):
Sampling interval: 50 μs (20 kHz)
Recording mode: Voltage Clamp
Number of sweeps: 10
DA: Send to Stim1 (send stimulus to amplifier 1), apply StimScale
AD: Imon-1 (A), Compression 1 (single sample, 2-byte integer)
Figure 6. Voltage pulses for leak subtraction. Left: For clarity, the voltage waveform corresponding to the three middle segments of the voltage stimulus in Figure 5 is redisplayed here (Vhold = -90 mV, 20 ms; -20 mV, 100 ms; Vhold, 100 ms). Right: The voltage waveform used as leak pulse stimulation, with the middle segment representing a scaled version of the middle segment in the main voltage stimulus at left (Vhold = -90 mV, 20 ms; -70 mV, 100 ms; Vhold, 100 ms).
FilterFactor: set to a value (≥2) that results in a lowpass filter cutoff frequency identical to that used for acquisition of the current evoked by the main stimulus
Number of segments: 3
Segment 1: constant, Vhold, 20 ms (duration identical to Segment 2 for the main stimulus)
Segment 2: constant, depolarization 20 mV relative to Vhold, should not be more depolarized than -60 mV, 100 ms (duration identical to Segment 3 for the main stimulus)
Segment 3: constant, Vhold, 100 ms (duration identical to Segment 4 for the main stimulus)
Note: The amplitude of the depolarization for Segment 2 should in principle be as large as possible (to optimize the signal-to-noise ratio), but not so large that it activates voltage-gated currents.
b. For performing the actual leak subtraction, it is a challenge that with data acquired using the voltage stimulus sequences described earlier (with different durations of the main and leak responses), the leak subtraction cannot be performed within the Patchmaster/Fitmaster environment. A working solution is to export the data and do the analysis in a different environment, e.g., IGOR Pro. In the example code provided below, it is assumed that wMain is the wave with the main response (potentially an average of ≥2 repetitions), and wLeak is the baseline-subtracted average of the leak responses.
Duplicate/O wMain, wMain_LS // wMain_LS will contain the leak-subtracted response
wMain_LS[A, B] -= wLeak(x) // A and B are the points corresponding to the start and
// end of Segment2 and Segment 4, respectively, of the
// main voltage stimulus
Alternatively, one can instead duplicate the part of wMain between points A and B and subtract wLeak from the copy:
Duplicate/O/R=[A, B] wMain, wMain_LS
wMain_LS -= wLeak
5. Settings for the LockIn Configuration window in Patchmaster (menu: Windows/LockIn).
Enter the following settings:
LockIn Mode: Sine + DC
Calibration Mode: Manual
Phase Shift: 0°
Attenuation: 1.000
[ ] Write to Notebook // optional, when hatched on, online analysis results will be printed
// to the Notebook window
Points to Average: Off
Generate Traces for: all amplifiers
Offline Computation - Traces to create:
[ × ] CM // membrane capacitance
[ × ] GM // membrane conductance
[ × ] GS // series conductance (inverse of Rseries)
[ × ] DC // conductance
Default Y-ranges:
Real (Y): 200 n Real (Z): 1.000
Imag (Y): 200 n Imag (Z): 1.000
Admit (Y): 200 nS Imp (Z): 1.000 Ω
Phase: 180.0°
CM: 40.00 pF DC: 4.000 nS
GM: 4.000 nS CV: 40.00 pF
GS: 400.0 nS GP: 400.0 nS
[ × ] V-rev: -15.00 mV [ ] Skip: 0
For an example of depolarization-evoked increase in capacitance (reflecting exocytosis) for an AII amacrine cell, see Figure 7.
Note: For EPC10 amplifiers with the "C-fast extended range" option, make sure that this feature is turned OFF before starting data acquisition.
Figure 7. Measuring depolarization-evoked exocytosis in an AII amacrine cell in a rat retinal slice. A. AII amacrine filled with Alexa 594 during whole-cell recording. Maximum intensity projection generated from widefield fluorescence image stack after deconvolution. Scale bar: 10 μm. B. Using the "Sine + DC" technique as implemented in the software LockIn amplifier implemented in Patchmaster software (with an EPC10 amplifier) to measure exocytosis-evoked capacitance increase in whole-cell recording of an AII amacrine cell (same cell as in A). Sine wave stimulation (2 kHz, ±15 mV from Vhold = -90 mV; top) before and after a 100 ms depolarization to 20 mV to activate voltage-gated Ca2+ channels and Ca2+-dependent exocytosis. Current responses evoked by the sine wave stimuli and depolarizing voltage pulse, displayed without (top) and with (bottom) leak subtraction. The inset shows Ca2+ current (with leak subtraction) at a higher time resolution. For each sine wave cycle, one data point was obtained for cell capacitance (Cm), cell membrane conductance (Gm), and series conductance (Gs). The resulting traces are displayed after baseline subtraction (ΔCm, ΔGm, and ΔGs). In the example illustrated here, leak subtraction was performed for the full duration of the acquired current. The depolarization-evoked increase of Cm (ΔCm = ~60 fF) was accompanied by an increase of Gs, but not by a change of Gm. Modified from ref. [18].
P. Capacitance measurement of endocytosis
In some cases, it might be of interest to measure endocytosis that typically follows exocytosis, although this process occurs at a much slower rate. Such measurements can be made by repeated application of a simple sine wave stimulus, e.g., a 2 kHz sine wave applied for 100 ms, repeated at an interval of, e.g., 0.5 s.
Q. Designing an experiment for capacitance measurement of exocytosis evoked by depolarization with arbitrary waveforms
Note: Designing the stimulus and acquisition sequence should be done before the day of the experiment, but for convenience, it is described here.
In addition to studying exocytosis evoked by Ca2+ influx through Cav channels opened by a standard square-wave voltage pulse, one might be interested in using arbitrary voltage waveforms, e.g., corresponding to excitatory postsynaptic potentials (EPSPs) evoked by activation of a presynaptic input [26] (Figure 8). In the case of the EPC10 amplifier (in combination with Patchmaster software), it is relatively straightforward to stimulate a cell with an arbitrary voltage waveform by using a stand-alone file template containing the selected waveform. The challenge, however, is that the cell must also be stimulated with sine wave waveforms for measuring the capacitance before and after the depolarizing stimulus. One solution is to stimulate the cell in parallel with two stimuli that when correctly configured simply get added to generate the final stimulus. With reference to the PGF editor window of Patchmaster, the core idea is to use two DA output channels. The first DA output channel is internally linked to the selected amplifier and used to apply the sine wave stimuli (configured to be output before and after the arbitrary waveform). The second DA output channel is for the arbitrary waveform, with the output sent to the EXTERNAL STIMULUS INPUT for voltage-clamp stimulation of the selected amplifier. When correctly configured, the sum of the DA outputs will stimulate the cell sequentially, first with a sine wave (for baseline capacitance measurement), then with the arbitrary waveform, and finally with a second sine wave (for post-stimulus capacitance measurement). Below is an example of how this can be configured. Notice that for this example, Vhold is set to -90 mV.
Figure 8. Exocytosis of an AII amacrine cell evoked by voltage-clamp depolarization with an excitatory postsynaptic potential (EPSP) waveform. A. Voltage stimulus applied to AII amacrine, with sine wave stimulation (2 kHz, ± 20 mV from Vhold = -90 mV) applied before and after depolarization with a stimulus waveform corresponding to an EPSP previously recorded in a different AII amacrine in response to depolarization of a presynaptic rod bipolar cell. B. Expanded view of voltage-clamp stimulus waveform in A. C. Current evoked in AII by sine wave stimulation and EPSP stimulus waveform (no leak subtraction). D. Total membrane capacitance (Cm) before and after depolarizing AII with EPSP waveform. The depolarization evoked a capacitance increase of ~50 fF. Modified from [26].
1. Configure the voltage stimulus as follows in the PGF editor of Patchmaster:
Sampling interval: 50 μs (20 kHz) // corresponds to fsine × 10
Recording mode: Voltage Clamp
DA Channel-1: Stim1, apply StimScale, use for LockIn
DA Channel-2: DA-6, use with FileTemplate
AD #1: Imon-1 (A), Compression 1 (single sample, 2-byte integer)
AD #2: Vmon-1 (V), Compression 1 (single sample, 2-byte integer)
AD #3: LockIn_CM (F), Compression 10 (single sample, 4-byte real)
AD #4: LockIn_GM (S), Compression 10 (single sample, 4-byte real)
AD #5: LockIn_GS (S), Compression 10 (single sample, 4-byte real)
FilterFactor: 5 (results in a lowpass filter cutoff frequency of 4 kHz = fsine × 2)
Number of segments: 7
DA Ch1:
Segment 1: constant, -90 mV, 100 ms
Segment 2: sine wave stimulus, -90 mV, 2000 ms
Segment 3: constant, -90 mV, 20 ms
Segment 4: constant, -90 mV, 180 ms // 180 ms is an example, the exact length depends on
// the specific waveform selected
Segment 5: constant, -90 mV, 100 ms // to let the membrane conductance return to baseline
Segment 6: sine wave stimulus, -90 mV, 2,000 ms
Segment 7: constant, -90 mV, 100 ms
DA Ch2: Segment 1 - Segment 7 all at 0 mV, with timing specification as for DA Ch1 (i.e., select Common timing).
2. Configure the sine wave parameters (for DA Ch1) as indicated below (in the Sinewave Parameters window):
"Use as LockIn SineWave"
Peak Ampl. [mV]: 20 (value) // "value" indicates that the numerical value can be modified
// online "Peak amplitude" is amplitude from baseline, the total
// amplitude (peak-to-peak) will be twice as large
Requested frequency: 2.0 kHz
Actual freq.: 2.0 kHz // may differ from "Requested freq."
Points / Cycle: 10
Cycles to Skip: 5 // setting to "5" discards the data points of the first five cycles to avoid
// "swing in" effects
Cycles to Average: 1 // setting to "1" provides one measurement point for each cycle
Total Cycles: 9002
V-reversal [mV]: -15.0 // the estimated value of the reversal potential of the leak current,
// the exact value is not very critical for the estimation of Cm, Rm,
// and Rs
3. Connect a BNC cable from DA-6 (output) to the input labeled EXTERNAL STIM. INPUT VC on the EPC10 amplifier (Amplifier-1).
4. During execution, make sure that the following settings are implemented (either manually or by incorporating them as statements in a Patchmaster protocol):
Vhold = -65 mV // the voltage at which the AII cell is held when not applying the stimulus
// of the PGF editor
External Stimulus Input = ON for Amplifier-1
External Scaling = 1.0 ×
5. Generate the (arbitrary) voltage stimulus waveform that will be used to depolarize the AII amacrine cell. In the example described below, we show how the waveform and file template can be generated using IGOR Pro. The example uses a waveform with an EPSP from a current-clamp recording of an AII amacrine cell. The EPSP was evoked by the depolarization of a rod bipolar cell recorded simultaneously with the AII amacrine cell. The first step is to duplicate the part of the recorded waveform ("voltRec") that contains the EPSP to a new waveform ("EPSP"). The first and last points of the segment of interest are arbitrarily set to 2025 and 3834, respectively, resulting in the EPSP waveform with 1810 points.
Duplicate/R=[2020,3834]/O voltRec, EPSP
The Y values at the beginning and end of this segment were -60.2 and -59.8 mV, respectively. The sampling interval of these waves was 0.1 ms (sampling frequency 10 kHz). For the new waveform to be used as a voltage-clamp stimulus template, we need a sampling interval of 0.05 ms (sampling frequency 20 kHz). Although the final voltage template is agnostic with respect to the sampling interval, the number of points needs to be correct. This can be achieved by interpolation of the waveform EPSP, generating a new waveform EPSPinterp:
Duplicate/O EPSP, EPSPinterp
Interpolate2/T=2/N=3620/E=2/Y=EPSPinterp EPSP
We need to make two constant waveforms and add them before ("PreSeg") and after ("PostSeg") EPSPinterp:
Make/O/N=42400 PreSeg // 2120 ms for an X interval of 0.05 ms (duration of Segments
// 1, 2, and 3)
Make/O/N=44000 PostSeg // 2200 ms for an X interval of 0.05 ms (duration of Segments
// 5, 6, and 7)
The voltage template stimulus will be output at Channel-2 (DA-6) and added to another voltage command (generated for Channel-1) with Vhold = -90 mV. Therefore, an offset must be added to the voltage waveform of the EPSP for the summed voltage to be identical to that of the EPSP waveform. This can be done by adding +90 mV to the EPSP waveform:
Duplicate/O EPSPinterp, EPSPinterp_offset
EPSPinterp_offset += 90e-3
Now the waveforms PreSeg, EPSPinterp_offset, and PostSeg can be concatenated:
Concatenate/NP/O {PreSeg, EPSPinterp_offset, PostSeg}, EPSPcmd
To avoid capacitive transients close to the EPSP waveforms, we still need to modify two short segments before (Segment 3, 20 ms) and after (Segment 5, 100 ms) the EPSP waveform itself (Segment 4) to make sure that the final voltage sums to approximately -60 mV (similar to that of the beginning and end of the EPSP waveform itself). Because the stimulus generated for Channel-1 sets Vhold = -90 mV, we need to set the voltage of these brief segments to +30 mV:
EPSPcmd[42000, 42399]=30e-3 // 399 points, 20 ms
EPSPcmd[46020, 48019]=30e-3 // 2000 points, 100 ms
The waveform can then be saved as a stimulus template file. The IGOR Pro procedure file PM_FileTemplate_v1.ipf contains code for this purpose. The name of the template file will automatically be provided with the suffix ".tpl” but must end with "_X" where "X" corresponds to the channel number (in the PGF editor window) where the file template will be used. In the example above, the file template is output on DA Channel-2 and the file name should be dCm_EPSPcmd_2.tpl. It must reside in a user-created folder named dCm_EPSPcmd that should reside directly inside the folder containing the Patchmaster application.
Apart from the special conditions used for data acquisition, other aspects of the recording are identical to those used for depolarizing cells with a square-wave pulse, including intra and extracellular solutions.
R. Designing an experiment for capacitance measurement of exocytosis evoked by depolarization via excitatory synaptic input
Note: Designing the stimulus should be done before the day of the experiment, but for convenience, it is described here.
In addition to the situation described in section Q above, where Ca2+ influx and subsequent exocytosis are driven via depolarization by an arbitrary waveform, it might also be of interest to measure capacitance in an experiment where an AII amacrine is depolarized by excitatory input from a presynaptic neuron, i.e., from a rod bipolar cell or an OFF-cone bipolar cell [26] (Figure 9). The design of this experiment is biologically more realistic, but also more challenging, for a number of reasons. First, the experiment requires simultaneous, dual recording of a bipolar cell and an AII amacrine cell. Second, for an EPSP to be generated in the AII amacrine, this cell must be in current clamp, not voltage clamp, when it receives the synaptic stimulus. Third, the measurement of capacitance immediately before and after the AII amacrine receives the synaptic input requires that the AII is in voltage clamp. Ideally, the instrumentation must permit rapid switching between voltage-clamp and current-clamp recording. Fourth, to mimic natural conditions as far as possible with respect to the ability of the synaptic input to evoke a normal EPSP, the AII amacrine should not be recorded with a Cs+-based intracellular solution. Instead, the solution must be K+-based, which will increase the noise for sine wave measurements of capacitance. Finally, to enable excitatory synaptic transmission, it is not possible to use pharmacological blockers of non-NMDA receptors, which will also contribute to an increase in noise during the recording. Nonetheless, these experiments are possible and provide useful information. For the experiment described here, the dual recording was performed for a rod bipolar cell and an AII amacrine cell. Notice that for this example, Vhold is set to -60 mV for both the AII amacrine and the rod bipolar cell.
Figure 9. Excitatory synaptic input from a rod bipolar (RB) cell evokes exocytosis in an AII amacrine cell. A. Synaptically coupled RB (magenta; cell filled with Alexa 594 during whole-cell recording) and AII amacrine (green; cell filled with Alexa 488 during whole-cell recording). Maximum intensity projection from widefield fluorescence image stacks. Same cell pair in A–C. Scale bar: 10 μm. B. Depolarization (200 ms) of RB (VRB) evoked EPSC in AII. As expected, depolarization of AII (VAII) evoked no response in RB. C. Exocytosis-evoked capacitance increase of AII triggered by EPSP evoked in AII by stimulation of RB. Magenta traces: RB voltage (VRB) and current (IRB). Green traces: AII voltage (VAII), current (IAII), and capacitance (Cm). During the first period, the AII was in voltage clamp (Vhold = -90 mV), and sine wave stimulation (2 kHz) was used to measure baseline capacitance (0–2.2 s). At 2.22 s, VAII was changed to -60 mV. After 50 ms, acquisition was temporarily halted, and AII was switched from voltage to current clamp (AII: VC → CC) to allow synaptic input from RB to evoke an EPSP. After resuming acquisition, RB was depolarized from -60 to -20 mV (100 ms duration; onset at ~2.84 s) to evoke EPSP in AII. At the end of the RB depolarization, AII was switched rapidly from current to voltage clamp (AII: CC → VC). After another 100 ms (to let the membrane conductance return to baseline), capacitance measurement was resumed. Note that AII capacitance increased by ~40 fF after the EPSP. Modified from [26].
1. Configure the setup with two micromanipulators, one for each cell. For the simultaneous, dual recording of a cell pair, the easiest way is to use an amplifier with two or more individual amplifier units, each with its own headstage. In the example illustrated below, an EPC10-USB-Quadro amplifier was used, with the AII amacrine cell recorded by Amplifier-1 and the rod bipolar cell recorded by Amplifier-3.
2. Establish dual whole-cell recording for a rod bipolar cell and an AII amacrine cell.
a. Find an AII amacrine cell as described above.
b. When a cell likely to be an AII has been found, see if a cell body in the distal part of the inner nuclear layer can be found, with location and shape typical of a rod bipolar cell. To increase the likelihood of finding a synaptically connected pair, the cell body of the putative rod bipolar cell should not be too far displaced laterally relative to the AII cell body (at most 1–2 cell body diameters).
c. Fill a pipette with intracellular pipette solution for the rod bipolar cell (IC4202), mount it in one of the pipette holders, and position the tip in the bath. Set the liquid junction potential to +14.5 mV.
d. Fill another pipette with intracellular pipette solution for the AII amacrine cell (IC4101), mount it in the other pipette holder, and position the tip in the bath. Set the liquid junction potential to +14.5 mV.
e. Find both pipettes under the objective and lower them such that they are located just above the surface of the slice.
f. Establish a GΩ-seal for the AII amacrine.
g. Establish a GΩ-seal for the rod bipolar cell.
h. Establish the whole-cell recording configuration for the AII amacrine cell. Capture the characteristic electrophysiological signature with action currents evoked by 5–10 mV depolarizing voltage pulses from -60 or -65 mV. This must be done quickly before the diffusion of QX314 from the pipette solution blocks the Nav channels in the AII.
i. Establish the whole-cell recording configuration for the rod bipolar cell.
3. Test for synaptic connectivity by applying a PGF sequence with two sweeps that together test for connectivity mediated by either chemical and/or electrical synapses in both directions (Figure 9B). For this, configure the voltage stimulus as follows in the PGF editor of Patchmaster:
Sampling interval: 100 μs (10 kHz)
Recording mode: Channel 1 - Voltage Clamp
Channel 2 - Voltage Clamp
Number of Sweeps: 2
DA Channel-1: Stim1, apply StimScale, relative to Vhold
DA Channel-3: Stim3, apply StimScale, relative to Vhold
AD #1: Imon-1 (A)
AD #2: Vmon-1 (V)
AD #3: Imon-3 (A)
AD #4: Vmon-3 (V)
FilterFactor: 5 (results in a lowpass filter cutoff frequency of 2 kHz)
Number of segments: 5
DA Ch1 (Stim1):
Segment 1: constant, Vhold, 200 ms
Segment 2: constant, -30 mV, 200 ms, V-incr. Mode: Increase, V-fact./incr. [mV]: +70 mV
Segment 3: constant, Vhold, 600 ms
Segment 4: constant, Vhold, 200 ms
Segment 5: constant, Vhold, 200 ms
DA Ch2 (Stim3): // select "Common timing"
Segment 1: constant, Vhold, 200 ms
Segment 2: constant, Vhold, 200 ms
Segment 3: constant, Vhold, 600 ms
Segment 4: constant, -30 mV, 200 ms, V-incr. Mode: Increase, V-fact./incr. [mV]: +70 mV
Segment 5: constant, Vhold, 200 ms
Note: If the pair consists of a rod bipolar cell presynaptic to an AII amacrine cell, this will be revealed by a synaptic response evoked by depolarization of the rod bipolar cell but no response in the rod bipolar cell by depolarization of the AII amacrine (Figure 9B). If a response is observed in the bipolar cell when the AII is depolarized, the bipolar cell is most likely an OFF-cone bipolar cell. If a postsynaptic response is evoked in either cell by hyperpolarization of the other cell (together with postsynaptic responses evoked by depolarization of either cell), the connectivity is likely to be mediated by electrical instead of chemical synapses, and the bipolar cell is most likely an ON-cone bipolar cell.
4. Assuming that the cell pair has been identified as a rod bipolar and an AII amacrine connected via a chemical synapse, apply the following PGF sequences and amplifier operations (must be configured before the experiment day, see point #5 below). Briefly, the fairly complex operation does the following:
1st PGF sequence (dCmABbaseline): AII in voltage clamp, apply sine wave stimulus to measure capacitance
Rod bipolar in voltage clamp, no stimulus
Change Vhold for AII to -60 mV and switch from voltage clamp to current clamp
2nd PGF sequence (dCmABepsp): Rod bipolar in voltage clamp, step to -20 mV to evoke exocytosis
Record EPSP in AII (in current clamp)
Change AII rapidly from current clamp to voltage clamp
AII in voltage clamp, apply sine wave stimulus to measure capacitance
Note: Use the Protocol Editor of Patchmaster to execute the various commands and operations in a tightly controlled and reproducible way. For additional documentation and examples of usage, see the manufacturer's website (http://www.heka.com/downloads/downloads_main.html#down_patchmaster). The EPC10 + Patchmaster system can apply the rapid mode switching (voltage clamp vs. current clamp) in both directions, but only a single time within a given PGF stimulus sequence. Accordingly, the series of operations indicated above has prioritized rapid switching (from current clamp to voltage clamp) following the synaptic stimulation of an AII amacrine by a rod bipolar cell.
5. Configure the PGF sequence dCmABbaseline as follows in the PGF editor of Patchmaster:
Sampling interval: 10 μs (100 kHz) // because the 2nd PGF makes use of the "rapid mode
// switching" during acquisition, the sampling frequency
// must be the highest possible
Recording mode: Channel 1 - Voltage Clamp // for AII amacrine cell
Channel 2 - Voltage Clamp // for rod bipolar cell
DA Channel-1: Stim1, apply StimScale, use for LockIn
DA Channel-2: Stim3, apply StimScale
AD #1: Imon-1 (A), Compression 1 (single sample, 2-byte integer)
AD #2: Vmon-1 (V), Compression 1 (single sample, 2-byte integer)
AD #3: LockIn_CM (F), Compression 100 (single sample, 4-byte real)
AD #4: LockIn_GM (S), Compression 100 (single sample, 4-byte real)
AD #5: LockIn_GS (S), Compression 100 (single sample, 4-byte real)
AD #6: Imon-3 (A), Compression 1 (single sample, 2-byte integer)
FilterFactor: 50 (results in a lowpass filter cutoff frequency of 2 kHz = fsine × 2)
Number of segments: 4
DA Ch1: // for AII amacrine cell
Segment 1: constant, -90 mV, 250 ms, start acquisition after 50 ms to avoid sampling capacitive transient
Segment 2: sine wave stimulus, -90 mV, 2000 ms
Segment 3: constant, -90 mV, 20 ms
Segment 4: constant, -65 mV, 50 ms
DA Ch2: // for rod bipolar cell
Segment 1 - Segment 4 all at Vhold, timing specification as for DA Ch1 ("Common timing").
6. Configure the sine wave parameters (for Segment 2 for DA Ch1) as indicated below (in the "Sinewave Parameters" window of Patchmaster):
"Use as LockIn SineWave"
Peak Ampl. [mV]: 20 (value) // "value" indicates that the numerical value can be modified
// online "Peak amplitude" is amplitude from baseline, the total
// amplitude (peak-to-peak) will be twice as large
Requested frequency: 1.0 kHz
Actual freq.: 1.0 kHz
Points/Cycle: 100
Cycles to Skip: 1
Cycles to Average: 1
Total Cycles: 2320
V-reversal [mV]: -60.0
7. Settings for the LockIn Configuration window in Patchmaster (for both dCmABbaseline and dCmABepsp PGF sequences).
a. Enter the following settings:
LockIn Mode: Sine + DC
Calibration Mode: Manual
Phase Shift: 0.0°
Attenuation: 1.000
Parent Trace: Linked Trace
[ ] Write to Notebook // optional, when hatched on, online analysis results will be printed
// to the Notebook window
Points to Average: Off
Generate Traces for: all amplifiers
Offline Computation - Traces to create:
[ × ] CM // membrane capacitance
[ × ] GM // membrane conductance
[ × ] GS // series conductance (inverse of Rseries)
[ × ] DC // conductance
Default Y-ranges:
Real (Y): 200 n Real (Z): 1.000
Imag (Y): 200 n Imag (Z): 1.000
Admit (Y): 200 nS Imp (Z): 1.000 Ω
Phase: 180.0°
CM: 40.00 pF DC: 4.000 nS
GM: 4.000 nS CV: 40.00 pF
GS: 400.0 nS GP: 400.0 nS
[ × ] V-rev: -60.00 mV [ ] Skip: 0
8. Amplifier control before, during, and after execution of the two PGF sequences
a. Before execution of the PGF sequence dCmABbaseline, make sure that the amplifier setting Gentle CC-Switch is set to ON. This means that when the amplifier is changed from voltage-clamp mode to current-clamp mode, the voltage-clamp holding current will be applied in current clamp to ensure that the membrane potential does not change appreciably.
b. After execution of dCmABbaseline, use the Protocol Editor to change Amplifier-1 from voltage clamp to current clamp.
c. Before execution of the PGF sequence dCmABepsp, change the amplifier setting for Gentle CC-Switch to OFF.
d. Execute the PGF sequence dCmABepsp.
9. Configure the voltage stimulus for the PGF sequence dCmABepsp as follows in the PGF editor of Patchmaster:
Sampling interval: 10 μs (100 kHz) // because the PGF makes use of the "rapid mode
// switching" during acquisition, the sampling frequency
// must be the highest possible
Recording mode: Channel 1: Any Mode // for AII amacrine cell
Channel 2: Voltage Clamp // for internal commands
Channel 3: Voltage Clamp // for rod bipolar cell
DA Channel-1: Stim1, apply StimScale, use for LockIn
DA Channel-2: Dig-out (word), absolute voltage // one output channel is set to Dig-out
// and used to send commands to the
// amplifier for the rapid mode switching
DA Channel-3:Stim3, apply StimScale
AD #1: Imon-1 (A), Compression 1 (single sample, 2-byte integer),
Build Instructions: "; VC-Switch=3" // command (between " ") to switch to voltage clamp
// at the start of segment 3
AD #2: Vmon-1 (V), Compression 1 (single sample, 2-byte integer)
AD #3: LockIn_CM (F), Compression 100 (single sample, 4-byte real)
AD #4: LockIn_GM (S), Compression 100 (single sample, 4-byte real)
AD #5: LockIn_GS (S), Compression 100 (single sample, 4-byte real)
AD #6: Imon-3 (A), Compression 1 (single sample, 2-byte integer)
FilterFactor: 50 (results in a lowpass filter cutoff frequency of 2 kHz = fsine × 2)
Number of segments: 5
DA Ch1: // for AII amacrine cell
Segment 1: constant, Vhold, 20 ms // effectively in current-clamp mode, inject Ihold
Segment 2: constant, Vhold, 100 ms // effectively in current-clamp mode, inject Ihold
Segment 3: constant, -90 mV, 100 ms // to let the membrane conductance return to
// baseline rapid switch to voltage clamp at
// beginning of segment 3
Segment 4: sine wave stimulus, -90 mV, 2,000 ms
Segment 5: constant, -90 mV, 100 ms
DA Ch2: // for Dig-out, internal use for EPC10 amplifier
Segment 1 - Segment 5 all constant, 0 V, timing specification as for DA Ch1 ("Common timing").
DA Ch3: // for rod bipolar cell
Segment 1: constant, Vhold, timing specification as for DA Ch1 ("Common timing")
Segment 2: constant, -20 mV, timing specification as for DA Ch1 ("Common timing")
Segment 3: constant, Vhold, timing specification as for DA Ch1 ("Common timing")
Segment 4: constant, Vhold, timing specification as for DA Ch1 ("Common timing")
Segment 5: constant, Vhold, timing specification as for DA Ch1 ("Common timing")
10. Configure the sine wave parameters (Segment 4 for DA Ch1 in dCmABepsp) as indicated for dCmABbaseline (see point 6 above).
S. Calibration of phase shift and attenuation of the EPC10 amplifier for "Sine + DC" lock-in" capacitance measurements of exocytosis
In general, in the whole-cell configuration, there is an upper limit for the sine wave frequency that should be used for "Sine + DC" lock-in capacitance measurements. This is the so-called "break frequency," given by Gillis [2] as:
f
b
=
1
/
2
×
π
×
C
m
/
G
s
+
G
m
(1)
where Cm is the membrane capacitance, Gs is the series conductance (inverse of series resistance Rs), and Gm is the membrane conductance. For the MC 10 model cell (electric circuit supplied with the EPC10 amplifier) in the whole-cell configuration, the value of fb is approximately 1.6 kHz. At higher sine wave frequencies, a greater proportion of the capacitive currents drops across Rs and a parallel (uncompensated) fraction of C-fast. For low sine wave frequencies, it is adequate to keep the values for phase shift and attenuation in the Patchmaster LockIn Configuration window at 0° and 1.00, respectively, as these values are reasonably good approximations, in the sense that the calculated calibration mode works well enough. For more precise measurements, however, it is necessary to calibrate the phase shift and attenuation of the instrumentation. Three different procedures are available according to the documentation provided by the manufacturer: calculated, measured, and manual. Please notice that it is never wrong to use the option for manual calibration, it just requires a bit more work on the part of the user. For details see:
http://www.heka.com/support/tutorials/tutorials_down/pm_tutorial.pdf // Patchmaster Tutorial
http://www.heka.com/downloads/software/manual/m_patchmaster.pdf // Patchmaster manual
1. Calculated calibration: This method leaves the calculation of the calibration results to the Patchmaster software and does not require any involvement by the user.
2. Measured calibration: This mode uses a resistor but is only valid for sine wave frequencies up to approximately 2 kHz. The procedure is described in the documentation provided by the manufacturer (see Patchmaster Tutorial referenced above). Measured calibration is essentially a manual calibration that uses the 10 MΩ resistor in the MC 10 model cell provided with the EPC10 amplifier. It is quite accurate for the range of sine wave frequencies where uncompensated stray capacitances do not play a significant role.
3. Manual calibration: For capacitance measurements with higher sine wave frequencies, it is better to use a capacitor than a resistor for the calibration. Either use a capacitor and read out the measurements of phase shift and attenuation, or, alternatively, use "capacitance dithering" (see Patchmaster Tutorial referenced above). Directly measuring the phase shift of the capacitor is the easiest method but requires the measurements to be performed in the medium gain range of the EPC10 amplifier. If the high gain range is used, the total amount of capacitance that can be measured before bringing the amplifier into saturation is very small (tens to at most a few hundred femtoFarads), and it is necessary to use capacitance dithering. This method is described in detail by the manufacturer (see Patchmaster Tutorial referenced above). In both cases, the estimated values for phase shift and attenuation can be entered directly in the Patchmaster LockIn Configuration window. The method using manual calibration and reading out the values for the phase shift and attenuation is described in the following, with example values from a real calibration session.
1. Manual calibration of phase shift and attenuation
a. Turn on the EPC10 amplifier and start the Patchmaster software. Wait ≥60 min for the amplifier to warm up before continuing.
b. Before performing a manual calibration of phase shift and attenuation, perform a standard calibration of the patch-clamp amplifier if needed (required approximately every 6 months). Consult the manufacturer's documentation for a step-by-step procedure.
2. Attach the MC 10 model cell to the headstage of the amplifier. Set the switch in the middle position (ON-CELL).
3. Generate a calibration voltage stimulus (PGF sequence) that applies a sine wave for capacitance measurement with identical sine wave frequency, filter settings, sampling rate, and gain as will be used for the subsequent real physiological measurements. There should be a sine wave segment with duration >100 ms (e.g., 500–1,000 ms).
4. Configure the LockIn Configuration window of Patchmaster.
a. Select ON-cell mode and Manual calibration.
b. Select (by checking) the option Write LockIn to Notebook.
c. Set the phase shift to 0° and the attenuation to 1.
d. Set the C-fast control field to 0.01 pF (i.e., lowest possible value) in the EPC10 amplifier window.
5. Execute the calibration voltage sequence.
a. The measured phase shift and capacitance will be written to the Notebook window.
b. Note the phase shift and subtract 90° since this is the magnitude introduced by the capacitor. The result corresponds to the phase shift of the instrumentation.
c. Enter this phase shift in the corresponding entry field of the LockIn Configuration window.
d. Keep the attenuation at 1.
6. Perform the "first" capacitance measurement.
a. Click the C-fast Auto button in the EPC10 amplifier window to compensate the (fast) capacitance.
b. Note the total capacitance in the C-fast field of the EPC10 amplifier window.
c. Execute the calibration voltage sequence again.
d. The capacitance written to the Notebook window should be very close to zero (typically just a few femtoFarads). This is the first measurement (zero capacitance, C-fast compensation).
7. Perform the "second" capacitance measurement.
a. Manually decompensate C-fast by a few picoFarads in the C-fast field of the EPC10 amplifier window (e.g., if the setting is 5.37 pF, change it to 3.37 pF).
b. Execute the calibration voltage sequence again and note the capacitance value. This is the second measurement.
c. The difference between the second and first measurements should in theory, with no attenuation, be equal to the magnitude of the magnitude of the C-fast decompensation. However, for high sine wave frequencies, this will not be the case, and the measured capacitance will be smaller than expected. This can be corrected by changing the Attenuation value in the LockIn Configuration window.
8. Tune the Attenuation value.
a. Essentially, the correction should be performed by reducing the setting for attenuation below 1 until the difference between the two measurements is equal to the magnitude of the decompensation.
b. The attenuation can also be calculated directly by dividing the measured difference by the magnitude of the decompensation.
c. Alternatively, one can perform a series of measurements for different values of decompensation and perform a linear curve fit of the graph that displays the measured difference versus the magnitude of the decompensation. The slope corresponds to the attenuation.
The calibrated values for phase shift and attenuation estimated with this technique will give improved results for capacitance measurements in the whole-cell configuration. For measurements using whole-cell recording, select this recording configuration in the EPC10 amplifier window and select Sine + DC and Manual calibration in the LockIn Configuration window.
9. Implement automatic adjustments when using multiple sine wave frequencies.
a. For experiments where several different sine wave frequencies are used, it becomes very cumbersome to manually change the values for phase shift and attenuation in the LockIn Configuration window.
b. Instead, the values can be sent from Patchmaster's Protocol Editor to the LockIn Configuration window with the following commands, which will set the LockIn parameters when the sine wave frequency changes during the execution of the experiment:
L LockInPS 330 // this will set the phase shift to 330°
L LockInAtt 0.9 // this will set the attenuation to 0.9
10. Example calibration
This section illustrates calibration results obtained for Amplifier-1 of an EPC10-triple (see Table 1 for typical results). In general, for a given sine wave frequency fc, the low-pass filter should be set to 2 × fc, and the filter factor should be set to 5. Accordingly, the sampling frequency (inverse of sampling interval) should be set to 5 × 2 × fc = 10 × fc. For the lower sine wave frequencies, the low-pass filter will correspond to Filter-2 of the EPC10, but the highest setting for this filter is 10 kHz. If the low-pass filter must be set higher than this, it is necessary to sample the output of Filter1, which has three different settings that can be used here (10, 30, and 100 kHz).
Table 1. Results from calibration of phase shift and attenuation for an EPC10 amplifier. Each value for the measured Cfast is the average of three repetitions. The values that must be provided to the LockIn Configuration window are the corrected phase shift in the third column (phase shift minus 90°) and the calculated attenuation in the sixth column (attenuation).
fsine (Hz) Phase shift (measured) Phase shift minus 90° Decompensated capacitance (pF) Measured Cfast (pF) Attenuation (Cmeasured/Cdecompensated)
100 10.3 -79.7 2 1.950 0.9750
200 8.7 -81.3 2 1.927 0.9637
400 7.2 -82.8 2 1.957 0.9785
1,000 357.4 267.4 2 1.919 0.9595
2,000 346.1 256.1 2 1.911 0.9555
4,000 343.8 253.8 2 1.795 0.8975
5,000 337.4 247.4 2 1.740 0.8702
10,000 325.9 235.9 2 1.335 0.6877
Data analysis
Here, we present a brief overview of the theory that forms the basis for the data analysis, with the goal of estimating Cm, Gm, and Gs from the current recorded in response to the sine wave voltage stimulus (for more advanced analysis, see [2]). As illustrated in Figure 10, when a time-varying voltage-clamp stimulus is applied to a round cell recorded in the whole-cell configuration of the patch-clamp technique, there are two pathways for the evoked current: a resistive current (IRm) will flow through the membrane resistance (Rm) and a capacitive current (ICm) will flow through the membrane capacitance (Cm; Figure 10A). For a sinusoidal stimulus, both IRm and ICm will be sinusoidal; whereas IRm will be in phase with the stimulus, ICm will be 90° phase shifted relative to the stimulus (Figure 10B). In addition, a third current component (IDC) will be generated if the (average) Vhold is different from the voltage source responsible for the resting membrane potential (Er in Figure 10A). The goal of a lock-in amplifier (implemented in hardware or software) is to extract the three components IRm, ICm, and IDC from the total current recorded via the patch pipette (Itot or Ipip; Figure 10B).
Figure 10. Equivalent electrical circuit of whole-cell recording from a round, unbranched cell and capacitance measurement of exocytosis using the Sine + DC technique. A. Equivalent electrical circuit for a whole-cell recording of a round, unbranched cell. Rs = series resistance; Cm = membrane capacitance; Rm = membrane resistance; Er = voltage source responsible for any DC current present at Vhold [2]. B. Capacitive and resistive (Ohmic) currents evoked in whole-cell, voltage-clamp recording (as in A) by stimulation with a sine wave voltage stimulus (Vsine) with a frequency of 1 kHz. The total current (Itot) recorded with the patch pipette (Ipip) is the sum of the resistive current IRm, which flows through Rm and in phase with Vsine, the capacitive current ICm, which flows through Cm and 90° phase shifted relative to Vsine, and the steady-state current IDC. The goal of the analysis is to separate Itot into different parts and use the results to estimate the circuit components as indicated in A.
First, it is necessary to determine the phase of the sine wave voltage stimulus. This is done by fitting the voltage waveform with the function,
V
(
t
)
=
A
×
sin
(
2
π
f
sin
e
t
+
α
)
+
V
h
o
l
d
(2)
where A is the amplitude, α is the phase (in radians), and 2πfsine is equivalent to the angular frequency (ω). Next, the current response recorded from the cell is separated into the real and imaginary components, in phase and 90° out of phase with the voltage stimulus, respectively, by fitting it with the function,
I
(
t
)
=
A
1
×
s
i
n
(
2
π
f
sin
e
t
+
α
)
+
A
2
×
c
o
s
(
2
π
f
sin
e
t
+
α
)
+
I
D
C
(3)
where A1 is the amplitude of the real component, A2 is the amplitude of the imaginary component, α is the phase [determined by fitting with eq. 2 for V(t)], and IDC is the amplitude of the steady-state current (Ihold). Cm, Rm, and Rs can then be calculated from A1, A2, and IDC, according to the following equations (stated as eq. 28 in [2]):
C
m
=
1
ω
B
(
A
2
+
B
2
-
A
G
t
)
2
(
A
-
G
t
)
2
+
B
2
(4)
R
m
=
1
G
t
(
A
-
G
t
)
2
+
B
2
A
2
+
B
2
-
A
G
t
(5)
R
s
=
A
-
G
t
A
2
+
B
2
-
A
G
t
(6)
where A and B are the amplitudes of the real (A1) and imaginary (A2) components (obtained from eq. 3) normalized to the amplitude of the voltage sine wave stimulus and Gt = IDC/(Vhold - Er) [2]. If the value of Er is known, it can be used. If it is not known, it has been reported that the calculation is not very sensitive to the exact value (see the Patchmaster Manual and Tutorial referenced above). We have found it adequate and reasonable to set the value of Vhold to -15 mV, corresponding to a certain level of depolarization of the resting membrane potential in AII amacrine cells recorded with an intracellular pipette solution containing Cs+ and TEA+ ions that block several types of K+ channels.
For data acquired as described in the current protocol, note that the analysis of Cm, Gm, and Gs is performed automatically by the Patchmaster/Fitmaster software, as the software has access to all the relevant information needed for the calculations corresponding to the equations stated above. Capacitance measurements as described here can be relatively noisy, and it is typically necessary to further process the analysis results to reduce noise. This can be done by averaging the results for several sine wave cycles (e.g., 10). In addition, it is possible to low-pass filter the time series of capacitance values (e.g., at 10 or 20 Hz, depending on noise level). Finally, it is possible to average the results of repeated measurements.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s) from our laboratory:
Hartveit et al. [18]. Capacitance measurement of dendritic exocytosis in an electrically coupled inhibitory interneuron: an experimental and computational study. Physiol. Rep. 7(15) e14186. doi.org/10.14814/phy2.14186 (Figure 1, panel B; Figure 4, panels C–G; Figure 5, panels C–F; Figure 6, panels B, C; Figure 7, panels A–D; Figure 8, panels B–E; Figure 9, panels D–G)
Hartveit et al. [26]. Dendritic morphology of an inhibitory retinal interneuron enables simultaneous local and global synaptic integration. J. Neurosci. 42(9): 1630–1647 (Figure 7, panels B, C, E–H).
See also the following publication that first used capacitance measurement of exocytosis from AII amacrine cells:
Balakrishnan et al. [13]. Synaptic vesicle exocytosis at the dendritic lobules of an inhibitory interneuron in the mammalian retina. Neuron 87(3): 563–575. doi: 10.1016/j.neuron.2015.07.016.
General notes and troubleshooting
General notes
1. Note that one can often observe changes not only of Cm but also of Gs and potentially also of Gm (cf. Figure 7B). This is to be expected when the Sine + DC capacitance measurement technique is applied to cells that display more complicated branching structures that cannot be modeled as simple RC circuits and where the stimulus-evoked change in capacitance occurs at a distance from the location of the recording electrode, as is the case for an AII amacrine cell recorded at the soma. These and related issues were examined in detail using computational modeling with realistic compartmental models of AII amacrine cells in a previous study from our laboratory [18].
2. In addition to the combination of an EPC10 amplifier and Patchmaster software, the more recent dPatch amplifier operated via SutterPatch software (Sutter Instrument) can also perform capacitance measurements using the "Sine + DC" technique. We have not yet tested capacitance measurements with this amplifier, but the implementation of SutterPatch promises to make, e.g., the handling of arbitrary stimulus waveforms considerably easier than the procedure described in the current protocol.
3. We have not tested other combinations of patch-clamp amplifiers and acquisition software, e.g., from Molecular Devices/Axon Instruments, and are therefore not able to comment on the potential ability to perform capacitance measurements using the "Sine + DC" technique. In cases where Patchmaster and the LockIn extension are used with amplifiers from companies other than HEKA Elektronik, the full "Sine + DC" technique can most likely not be used. The alternative “Piecewise Linear” technique makes no attempt to determine the actual value of the admittance or any of the three equivalent circuit parameters. Instead, the measured change in admittance is used to estimate changes in the relevant parameters. For additional details, see the documentation from HEKA Elektronik and [2].
4. For advice on the construction of a setup for patch-clamp recording, see the detailed description by Penner in [27].
Troubleshooting
Problem 1: Series resistance is unacceptably high (e.g., ≥ 50 MΩ).
Possible cause: Patch pipettes are too small (the opening of the distal tip is too small).
Solution: Pull new pipettes with lower resistance.
Problem 2: Cells swell or shrink (markedly) during recording.
Possible cause: The intracellular pipette solution has too low or too high osmolality.
Solution: Measure the osmolality of the pipette solution. Make sure to calibrate the osmometer first. Make new solutions if necessary.
Problem 3: Too many cells in the retinal slices do not look healthy.
Possible cause: Rat suffered from hypoxia during the procedure with inhalation anesthesia. Healthy cells should appear smooth in the microscope. When approached with a patch pipette, it is easy to make a dimple in the soft membrane. Unhealthy or sick cells can be recognized by a high-contrast and rough appearance [28,29].
Solution: Review the procedure for anesthetizing and killing the animal. Make sure that the rat is allowed to breathe 100% O2 for several minutes before being exposed to anesthetic inhalation.
Problem 4: Too many cells in the retinal slices do not look healthy (see Problem 3).
Possible cause: The buffered extracellular solution used for dissection was not prepared correctly or has been contaminated by microorganisms (visible as turbidity and/or color change of the solution).
Solution: Make new stock solutions (KCl, MgCl2, CaCl2) and buffer solution.
Problem 5: Too many cells in the retinal slices do not look healthy (see Problem 3).
Possible cause: The buffered extracellular solution used for bath perfusion was not made correctly.
Solution: Review the procedure for making the solution and adding the various chemicals. Make sure that CaCl2 is not added before the solution is equilibrated with 5% CO2. Make new stock solutions (KCl, MgCl2, CaCl2) and bath solution.
Problem 6: Too many cells in the retinal slices do not look healthy (see Problem 3).
Possible cause: The dissection technique is suboptimal or too slow.
Solution: Review the procedure for dissecting and handling the retinal tissue. Remember to handle the tissue very gently; it does not tolerate rough mechanical manipulation.
Problem 7: During dissection, the eye cup looks abnormal with one or more regions with discolored (often white) retinal tissue.
Possible cause: The animal might have been ill or is too old.
Solution: Check the status with the animal facility. Make sure that rats are only used up to a maximum age of 7–8 weeks (preferably not more than 6 weeks old).
Problem 8: Exocytosis runs down too fast.
Possible cause: Be aware that rundown is relatively fast and cannot be expected to last more than approximately 12–15 min.
Solution: Review general procedures as described above. Make fresh intracellular pipette solution.
Problem 9: The AII amacrine cell does not display the characteristic unclamped action current when stimulated with a depolarizing test pulse (5–10 mV depolarization) after breaking into the cell.
Possible cause: The cell is either not an AII amacrine cell or has lost its axon initial segment-like process during the procedure for making slices.
Solution: Try to find a new cell. If the problem occurs frequently, you may be targeting cells located too close to the surface of the slice. Try to target cells located at a deeper level in the slice (≥10 μm below the surface).
Problem 10: During simultaneous, dual recording of pairs of rod bipolar cells and AII amacrine cells, the cells are not synaptically connected.
Possible cause: Most likely, cells are located too far apart, with no possibility for synaptic contact.
Solution: Use fluorescence microscopy to visualize both cells and see if there is evidence for morphological contact. If not, try to find a rod bipolar cell with less lateral displacement relative to the cell body of the AII amacrine cell.
Problem 11: There is excessive noise during the recording.
Possible causes: AgCl coating of ground electrodes (one in the pipette, one in the bath) is too old or insufficient. The grounding of the rig/setup involves one or more ground loops.
Solution: First, make sure that the Ag-wires of both ground electrodes have been freshly chlorided (section D). Review and potentially rewire the ground connections of the setup.
Problem 12: It is not possible to obtain GΩ-seals.
Possible cause: There can be several reasons for this, including the deteriorating health of cells/retinal tissue and bad intracellular pipette solution.
Solution: Make sure that patch pipettes are freshly pulled and have clean tips (observe under a microscope). Change to a vial with fresh intracellular pipette solution.
Supplementary information
The following supporting information can be downloaded here:
1. IGOR Pro procedure file PM_FileTemplate_v1.ipf
Acknowledgments
This protocol was adapted from Hartveit et al. [18] and Hartveit et al. [26]. This work was supported by the Trond Mohn Foundation (TMF) and the Mohn Research Center for the Brain. Financial support from The Research Council of Norway is gratefully acknowledged (NFR 182743, 189662, 214216 to E.H.; NFR 213776, 261914 to M.L.V.). We thank Áurea Castilho for excellent technical assistance.
Competing interests
The authors declare no competing interests.
Ethical considerations
The use of animals in the studies from our laboratory cited in this protocol was conducted under the approval of and in accordance with the regulations of the Animal Laboratory Facility at the Faculty of Medicine at the University of Bergen (accredited by AAALAC International).
References
Südhof, T. C. and Rizo, J. (2011). Synaptic Vesicle Exocytosis. Cold Spring Harbor Perspect Biol. 3(12): a005637–a005637.
Gillis, K. D. (1995). Techniques for Membrane Capacitance Measurements. In: Sakmann, B. and Neher, E. (Ed.). Single-Channel Recording. pp. 155–198. Plenum Press, New York, London.
Lindau, M. and Neher, E. (1988). Patch-clamp techniques for time-resolved capacitance measurements in single cells. Pflügers Arch. 411(2): 137–146.
Kim, M. and Von Gersdorff, H. (2010). Extending the realm of membrane capacitance measurements to nerve terminals with complex morphologies. J Physiol. 588(12): 2011–2012.
Kushmerick, C. and von Gersdorff, H. (2003). Exo-endocytosis at mossy fiber terminals: Toward capacitance measurements in cells with arbitrary geometry. Proc Natl Acad Sci USA. 100(15): 8618–8620.
Hallermann, S., Pawlu, C., Jonas, P. and Heckmann, M. (2003). A large pool of releasable vesicles in a cortical glutamatergic synapse. Proc Natl Acad Sci USA. 100(15): 8975–8980.
Heidelberger, R., Heinemann, C., Neher, E. and Matthews, G. (1994). Calcium dependence of the rate of exocytosis in a synaptic terminal. Nature. 371(6497): 513–515.
Oltedal, L. and Hartveit, E. (2010). Transient release kinetics of rod bipolar cells revealed by capacitance measurement of exocytosis from axon terminals in rat retinal slices. J Physiol. 588(9): 1469–1487.
Sun, J. Y. and Wu, L. G. (2001). Fast Kinetics of Exocytosis Revealed by Simultaneous Measurements of Presynaptic Capacitance and Postsynaptic Currents at a Central Synapse. Neuron. 30(1): 171–182.
Wölfel, M. and Schneggenburger, R. (2003). Presynaptic Capacitance Measurements and Ca2+ Uncaging Reveal Submillisecond Exocytosis Kinetics and Characterize the Ca2+ Sensitivity of Vesicle Pool Depletion at a Fast CNS Synapse. J Neurosci. 23(18): 7059–7068.
Hsu, S. F. and Jackson, M. B. (1996). Rapid exocytosis and endocytosis in nerve terminals of the rat posterior pituitary. J Physiol. 494(2): 539–553.
Zhou, Z. Y., Wan, Q. F., Thakur, P. and Heidelberger, R. (2006). Capacitance Measurements in the Mouse Rod Bipolar Cell Identify a Pool of Releasable Synaptic Vesicles. J Neurophysiol. 96(5): 2539–2548.
Balakrishnan, V., Puthussery, T., Kim, M. H., Taylor, W. R. and von Gersdorff, H. (2015). Synaptic Vesicle Exocytosis at the Dendritic Lobules of an Inhibitory Interneuron in the Mammalian Retina. Neuron. 87(3): 563–575.
Sassoè-Pognetto, M., Wassle, H. and Grunert, U. (1994). Glycinergic synapses in the rod pathway of the rat retina: cone bipolar cells express the alpha 1 subunit of the glycine receptor. J Neurosci. 14(8): 5131–5146.
Veruki, M. L. and Hartveit, E. (2002). AII (Rod) Amacrine Cells Form a Network of Electrically Coupled Interneurons in the Mammalian Retina. Neuron. 33(6): 935–946.
Veruki, M. L. and Hartveit, E. (2002). Electrical Synapses Mediate Signal Transmission in the Rod Pathway of the Mammalian Retina. J Neurosci. 22(24): 10558–10566.
Zandt, B. J., Veruki, M. L. and Hartveit, E. (2018). Electrotonic signal processing in AII amacrine cells: compartmental models and passive membrane properties for a gap junction-coupled retinal neuron. Brain Struct Funct. 223(7): 3383–3410.
Hartveit, E., Veruki, M. L. and Zandt, B. (2019). Capacitance measurement of dendritic exocytosis in an electrically coupled inhibitory retinal interneuron: an experimental and computational study. Physiol Rep. 7(15): e14186.
Neher, E. (1992). [6] Correction for liquid junction potentials in patch clamp experiments. In: Rudy, B. and Iverson, L. E. (Ed.). Methods in Enzymology “Ion Channels”. (vol. 207, p. 123–131). San Diego, CA: Academic.
Neher, E. (1995). Voltage Offsets in Patch-Clamp Experiments. In: Sakmann, B. and Neher, E. (Ed.). Single-Channel Recording. pp.147–153. Plenum Press, New York, London.
Mørkve, S. H., Veruki, M. L. and Hartveit, E. (2002). Functional characteristics of non‐NMDA‐type ionotropic glutamate receptor channels in AII amacrine cells in rat retina. J Physiol. 542(1): 147–165.
Heidelberger, R., Zhou, Z. Y. and Matthews, G. (2002). Multiple Components of Membrane Retrieval in Synaptic Terminals Revealed by Changes in Hydrostatic Pressure. J Neurophysiol. 88(5): 2509–2517.
Hull, C. and von Gersdorff, H. (2004). Fast Endocytosis Is Inhibited by GABA-Mediated Chloride Influx at a Presynaptic Terminal. Neuron. 44(3): 469–482.
Habermann, C. J., O'Brien, B. J., Wässle, H. and Protti, D. A. (2003). AII Amacrine Cells Express L-Type Calcium Channels at Their Output Synapses. J Neurosci. 23(17): 6904–6913.
Heinemann, S. H. (1995). Guide to Data Acquisition and Analysis. In: Sakmann, B. and Neher, E. (Ed.). Single-Channel Recording. pp. 53–91. Plenum Press, New York, London.
Hartveit, E., Veruki, M. L. and Zandt, B. J. (2022). Dendritic Morphology of an Inhibitory Retinal Interneuron Enables Simultaneous Local and Global Synaptic Integration. J Neurosci. 42(9): 1630–1647.
Penner, R. (1995). A Practical Guide to Patch Clamping. In: Sakmann, B. and Neher, E. (Ed.). Single-Channel Recording. pp. 3–30. Plenum Press, New York, London.
Sakmann, B. and Stuart, G. S. (1995). Patch-Pipette Recordings from the Soma, Dendrites, and Axon of Neurons in Brain Slices. In: Sakmann, B. and Neher, E. (Ed.). Single-Channel Recording. pp. 199–211. Plenum Press, New York, London.
Edwards, F. A. and Konnerth, A (1992). [13] Patch-clamping cells in sliced tissue preparations. In: Rudy, B. and Iverson, L. E. (Ed.). Methods in Enzymology “Ion Channels”. (vol. 207, p. 208–222). San Diego, CA: Academic.
Article Information
Publication history
Received: Aug 19, 2024
Accepted: Oct 25, 2024
Available online: Nov 14, 2024
Published: Jan 5, 2025
Copyright
© 2025 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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Automatic Adaptive Algorithm for Delineation of Cerebral-Spinal Fluid Regions for Non-contrast Magnetic Resonance Imaging Volumetry and Cisternography in Mice
RG Ryszard S. Gomolka
Published: Vol 15, Iss 1, Jan 5, 2025
DOI: 10.21769/BioProtoc.5148 Views: 189
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Original Research Article:
The authors used this protocol in eLIFE Feb 2023
Abstract
Magnetic resonance imaging (MRI) is an invaluable method of choice for anatomical and functional in vivo imaging of the brain. Still, accurate delineation of the brain structures remains a crucial task of MR image evaluation. This study presents a novel analytical algorithm developed in MATLAB for the automatic segmentation of cerebrospinal fluid (CSF) spaces in preclinical non-contrast MR images of the mouse brain. The algorithm employs adaptive thresholding and region growing to accurately and repeatably delineate CSF space regions in 3D constructive interference steady-state (3D-CISS) images acquired using a 9.4 Tesla MR system and a cryogenically cooled transmit/receive resonator. Key steps include computing a bounding box enclosing the brain parenchyma in three dimensions, applying an adaptive intensity threshold, and refining CSF regions independently in sagittal, axial, and coronal planes. In its original application, the algorithm provided objective and repeatable delineation of CSF regions in 3D-CISS images of sub-optimal signal-to-noise ratio, acquired with (33 μm)3 isometric voxel dimensions. It allowed revealing subtle differences in CSF volumes between aquaporin-4-null and wild-type littermate mice, showing robustness and reliability. Despite the increasing use of artificial neural networks in image analysis, this analytical approach provides robustness, especially when the dataset is insufficiently small and limited for training the network. By adjusting parameters, the algorithm is flexible for application in segmenting other types of anatomical structures or other types of 3D images. This automated method significantly reduces the time and effort compared to manual segmentation and offers higher repeatability, making it a valuable tool for preclinical and potentially clinical MRI applications.
Key features
• This protocol presents a fully automatic adaptive algorithm for the delineation of CSF space regions in 3D-CISS in vivo images of the mouse brain.
• The algorithm represents an analytical method for adaptive CSF regions separation based on cumulative distribution of brain image intensities and contrast calculation-based slice-wise region growing.
• Users can interactively alter the input parameters to modify the algorithm’s output in a variety of 3D brain MR and μCT or CT images.
• The algorithm is implemented in MATLAB 2021a and is compatible with all versions up to 2024a.
Keywords: MRI 3D-CISS CSF space volumetry Automatic 3D image segmentation
Graphical overview
From acquisition to automatically segmented CSF space image: a graphical abstract for application of the presented automatic adaptive algorithm for objective cerebrospinal fluid (CSF) space segmentation in 3D constructive interference steady-state (3D-CISS) images of the mouse brain.
Background
Magnetic resonance imaging (MRI) is an invaluable technique for anatomical and functional whole-brain in vivo imaging [1–3]. For non-invasive whole-brain MRI of the brain cerebrospinal fluid (CSF) space, a state-of-the-art T2-weighted spin-echo (SE) imaging can be used with inversion-recovery or saturation-recovery preparation pulses [4]. This provides high accuracy for the CSF space differentiation by means of “pure” image contrast [5], however, it requires remarkably long acquisition times reducing applicability, especially for 3D whole-brain imaging. An alternative considers rapid gradient-echo imaging based on steady-state techniques [6,7], encompassing radiofrequency-spoiled [8] gradient-echo (GRE) [9], to visualize high-intensity CSF regions. While providing shorter acquisition times than SE, GRE is strongly vulnerable to field inhomogeneities [10,11] requiring transmit field corrections [12]. Alternatively, strong T2 weighted contrast imaging with a true balanced steady-state free precession (TrueFISP) [13,14] allows higher signal-to-noise ratio (SNR) efficiency resulting in high spatial 3D resolution. TrueFISP is characterized with the image contrast close to that of SE [15], while for the time-to-repetition (TR) to time-to-echo (TE) ratio of 2:1 provides high T2 contrast [16–18]. It is easily affected by phase shift errors (banding artifacts) as it acquires constant phases across voxels [16,19,20]. A solution for CSF space volumetry uses maximum intensity projection of at least two combined TrueFISP acquisitions known as a 3D constructive interference steady-state sequence (3D-CISS), employed to visualize the structures at the skull base [21]. 3D-CISS provides strong T2-weighted CSF contrast and compensation for banding artifacts [22] and, due to balanced gradients, is almost invulnerable to flow artifacts (see Figure S1 for the mouse brain volume and CSF space visualization using 3D-CISS). This results in homogenous intensity distribution and high SNR and CSF space contrast [16–18,22,23]. Nevertheless, accurate delineation of the CSF space (similar to the segmentation of other structures in complex brain anatomy) is a critical part of MR image processing and analysis [24,25], especially when no external contrast agents are used for CSF space imaging. An accurate and reliable automatic approach is highly desired and possesses a clear advantage over the manual approach due to the time and effort required and higher repeatability. In the last decade, many segmentation methods for MR brain images in clinical [26–28] and preclinical [29–33] settings have been proposed, with a clear trend toward the application of artificial neuronal networks. However, in the vast majority of limited data available on-site, the analytical approach is fully sufficient. Recently, we have employed 3D-CISS along with a dedicated automatic adaptive algorithm in MATLAB for the objective delineation of CSF in aquaporin-4-null and wild-type mice [34]. Herein, we describe in detail this in-house developed algorithm, which provides automatic delineation of CSF space regions. The algorithm is based on the original method to study the cumulative distribution of intensities and contrast of the brain stroke regions in non-contrast computed tomography, characterized by similar brain tissue contrast and image SNR as in CISS [35,36], and provides objective delineation of the CSF space. The method is not limited to the purpose, and with input parameter adjustment, it can be used for segmentation of structures occupying different parts of voxel intensity distribution in the images.
Equipment
3D-CISS image acquisition
1. 9.4 Tesla animal scanner (Bruker BioSpin, Ettlingen, Germany, model: BioSpec 94/30 USR) or another preclinical scanner
2. 240 mT/m gradient coil (Bruker BioSpin, model: BGA-12S) or other
3. Cryogenically cooled transmit/receive (Tx/Rx) quadrature-resonator (Bruker BioSpin, model: CryoProbe) or other preferably volumetric Tx/Rx resonator providing sufficient sensitivity
Automatic CSF space delineation
1. Standard PC with > 2 cores processor and > 8 GB RAM memory (herein, Intel Core i7-10700U, 32 GB RAM)
Software and datasets
1. ITK-SNAP v3.x or later (www.itksnap.org, accessed July 2024) [37]
2. Analysis of Functional NeuroImages (AFNI; https://afni.nimh.nih.gov, accessed July 2024) [38]
3. FMRIB Software Library (FSL) v5.0 or later (https://fsl.fmrib.ox.ac.uk, accessed July 2024) [39]
4. MATLAB R2019a or later (https://www.mathworks.com/products/matlab.html, accessed July 2024). Current implementation developed in MATLAB R2021a, and tested in versions R2022b, R2023b, and R2024a.
5. All data code is available via GitHub (https://github.com/RSG1UCPH/CSF-space-segmentation-in-3D-CISS.git)
Procedure
A. Magnetic resonance imaging
The described algorithm was originally employed for objective assessment of the volumes and structural differences between the brain CSF spaces in aquaporin-4-null and wild-type littermate mice, based on 3D-CISS images [34]. MR imaging was performed at 9.4 Tesla (BioSpec 94/30 USR, Bruker BioSpin, Ettlingen, Germany) equipped with 240 mT/m gradient coil (BGA-12S, Bruker BioSpin) and cryogenically cooled transmit/receive (Tx/Rx) quadrature-resonator (CryoProbe, Bruker BioSpin). The imaging protocol consisted of T2-weighted 2D rapid acquisition with relaxation enhancement (RARE) for reference spatial planning and two 3D-TrueFISP acquisitions with orthogonal phase encoding directions for CSF space imaging. For details of the imaging protocol, please refer to the original paper [34].
B. MATLAB and toolboxes
MATLAB can be installed from the MathWorks product webpage and is available for Windows, Linux, and Mac operating systems (OS). For our purposes, a Windows OS was used. Additional information regarding available MATLAB products and installation on different OS can be found on the producer’s web page (www.mathworks.com/products/matlab.html, accessed July 2024).
The following MATLAB toolboxes are required to run the algorithm:
A. Image Processing Toolbox (functions: niftiread, bwboundaries, medfilt3, permute, and imshow)
B. Statistics and Machine Learning Toolbox (function: prctile)
C. Step-by-step protocol description
Note: Skip steps C1–3 if a 3D-CISS image is already available.
1. Image acquisition [34]: Acquire two 3D-TrueFISP volumes, each with opposite phase encoding direction (i.e., 0° and 180°).
2. Save acquired images in NIfTI format.
3. 3D-CISS image formation:
a. Perform motion-correction of 3D-TrueFISP images (if more than one repetition was used per each acquisition) 10 times or until no further improvement [3Dvolreg() function in AFNI], aiming to reduce the influence of random motion on the subsequently computed 3D-CISS image.
b. Calculate averaged 3D-TrueFISP image from all repetitions per acquisition using opposite phase encoding direction.
c. Realign the second averaged (and subsequent if available) 3D-TrueFISP volume (i.e., acquired with 180° phase encoding direction) to the first volume (i.e., acquired with 0° phase encoding direction) using rigid-body registration [6 d.f.; 3Dvolreg() function in AFNI] for subsequent calculation of 3D-CISS volume.
d. Calculate 3D-CISS image as a maximum intensity projection from two co-registered 3D-TrueFISP volumes, allowing to obtain an almost banding artifacts-free image (see example of acquired 3D-CISS image in Figure S2A).
4. Perform semi-automatic segmentation of brain parenchyma (Figure S2B) in calculated 3D-CISS image using the Segment 3D tool in ITK-SNAP or other semi-automatic or automatic method or software available for this purpose. Brain parenchyma is considered here as the brain tissue volume surrounded by dark regions of the skull image and including intracerebral vessels, as defined in [34] (see example in Figure S2C–E). This step is necessary to reduce the computational burden on subsequent CSF space segmentation in MATLAB, by removing the regions outside the brain parenchyma image from further analysis.
5. To correct for intensity inhomogeneities coming from the B0 field and the profile of the Tx/Rx coil (in original method surface profile of the CryoProbe [34]), perform N4 bias field correction in the brain-extracted parenchyma using FAST tool in FSL (0.5 sigma, 20 mm FWHM, 4 iterations).
6. Finally, perform automatic CSF space segmentation using the dedicated adaptive algorithm described below in MATLAB. For a single, bias-corrected, and brain-extracted 3D-CISS volume, the algorithm separates the ventricular and perivascular CSF spaces from the brain parenchyma image in three dimensions, and by means of four consecutive steps based on the following functions:
• CSF_volumetry(): The main function that runs the entire processing pipeline.
• mask_brain_only(): Isolates the brain regions from the image.
• intensify_CSF(): Enhances the CSF space image seed regions based on an adaptive thresholding.
• grow_regions(): Refines the seed regions by comparing their intensity and contrast to that of surrounding brain parenchyma, in sagittal plane.
• grow_regions2(): Further refines the fine-tuned seed regions from the previous step, in axial and coronal planes.
• SNRstats(): Computes basic statistical properties of the input image.
• save_nifti(): Function for saving the results of the automatic segmentation algorithm.
Note: The algorithm is available online (https://github.com/RSG1UCPH/CSF-space-segmentation-in-3D-CISS.git) and is equipped with a simplified user interface (see Figure 1A and B, function CSF_volumetry_gui()), allowing modification of the most important input parameters and analyzing a single image at a time. The CSF_volumetry() function can be used without the user interface and can be adapted for analyzing multiple files consecutively in a loop. For supplementary instructions on the usage of the functions, please refer to the README.txt file available along with the original code in MATLAB.
Figure 1. Action steps of the presented automatic adaptive algorithm for delineation of cerebrospinal fluid (CSF) space in non-contrast 3D constructive interference steady-state (3D-CISS) images of the mouse brain. (A) User interface for the CSF segmentation algorithm, where parameters such as plotting results, applying median filtration, and adjusting α and α2 (see step 4b) and percentile values (see steps 2–4) can be done before selecting a file and running the processing. (B) After the image is selected, its mid-slice is displayed in the window on the right side from the user interface input text boxes. (C) Brain parenchyma is enclosed within a calculated bounding box for the removal of non-continuous and not adjacent to the parenchyma image high-intensity regions (see step 1). Parenchymal regions are highlighted in each analyzed slice, with different outlines indicating areas of interest, namely: red outline, brain parenchyma bounding box (BOX, see step 1); green outline, bounding box covering only the olfactory and optic nerves area (distal 25% length of the red box + arbitrary, set to 16 voxels, extension outside the red box to cover any regions potentially removed by the red box); pink outline, refined brain parenchyma boundaries within the green box outline; blue outline, final brain parenchyma outline after removal of high-intensity parenchyma image regions not adjacent to the brain (BOX2, see step 1). (D) Binary mask image showing the segmented CSF seed regions before and (E) after optional 3D median filtration (see steps 2 and 3). The CSF seed regions mask further undergoes region growing in (F) sagittal, (G) axial, and (H) coronal planes, considering regions dilation (red outline) followed by erosion (green line; see step 4). (I) Confirmation dialog indicates successful completion of the CSF space segmentation, signaling the end of the automated analysis.
D. CSF space segmentation in an input 3D-CISS image
The CSF space segmentation in an input 3D-CISS image takes place in the following steps:
1. Computation of the bounding box enclosing the brain and removal of high-intensity regions not adjacent to the brain parenchyma as branches of the optic nerve’s residual after the semi-automatic brain image extraction.
The bounding box is automatically computed based on both minimization and maximization of the voxel intensity variance slice-wise, separately in three orthogonal planes. The parenchyma volume surrounded by the bounding box is being enclosed, and the solitary regions are removed based on their geometrical properties calculated slice-wise in the sagittal plane: eccentricity ≥ 0.5, roundness ≥ 0.5, perimeter < 0.005% of the brain parenchyma voxels count. The resulting brain image mask is geometrically dilated with a disk kernel of 11 pixels in diameter (Figure 1C) to enclose potentially removed or non-continuous parenchymal regions. The considered non-continuity appears in the case of residuals from banding artifacts at the borders of the skull and the ethmoidal bone. Subsequently, brain parenchyma image volume is updated according to the resulting mask for further automatic segmentation of the CSF space.
Important: This method assumes that the brain-extracted and bias-corrected brain parenchyma image is realigned to orthogonal axes and placed in the center of the 3D-CISS image (see example file: Test_brain1.nii.gz and Test_brain2.nii.gz).
Function Definition and Description
The function [BOX] = mask_brain_only(image, plt) isolates the brain region in a 3D volume by creating a binary mask that excludes non-brain areas.
Inputs: image (3D image volume) and plt (flag for plotting intermediate results).
Output: BOX (binary 3D mask for the brain image).
a. Middle slice calculation
The middle slice in the sagittal plane is found by averaging two central slices of the 3D-CISS volume. This helps to identify the central region of the brain and to subsequently calculate the extents of the bounding box. The mid-slice in the coronal plane is calculated after spatial coordinates of the bounding box are calculated in sagittal and axial planes.
b. Bounding box calculation
For calculation of the bounding box enclosing the brain parenchyma image in three dimensions, a variance is calculated across columns and rows of the mid-sagittal slice to identify brain parenchyma extents (with non-zero variance). Based on minimization and maximization of the calculated non-zero variance in sagittal, axial, and coronal planes, coordinates defining the bounding box are determined.
c. Binary brain mask
Two binary masks (BOX and BOX2) are initialized. BOX is the main mask, and BOX2 is a secondary mask to help refine the 3D brain-encapsulated volume. For each slice, the masks are refined by:
i. Removing areas with significant intensity.
ii. Dilating the mask to fill any gaps and smooth boundaries.
iii. Selecting regions based on shape properties (circularity, eccentricity, perimeter).
iv. Eroding the mask and filling holes.
The main mask (BOX) and the refined secondary mask (BOX2) are combined to produce the final brain region mask.
d. Applying the bounding box
The refined mask of the bounding box enclosing the brain parenchyma image in three dimensions is applied to the original image, and the result is saved into new_image 3D volume.
e. Plotting
If plotting is enabled, intermediate results are displayed to visualize the mask refinement process.
Note: The bounding box calculation is an integral part of the described method, and its omission is generally not recommended. However, users who thoroughly understand the method and code in MATLAB and are confident that the brain parenchyma is well separated in their input image may choose to bypass this step by commenting relevant fragment of the original code. Due to this possibility, the updated with the bounding box new_image, originally defined in the MATLAB code, is defined in the following functions as the image.
Important: The bounding box calculation assumes that, in 2D slice of the brain image volume, the olfactory bulb region is placed toward the end of the image, considering [0,0] point as the left-upper origin of the slice (see example file: Test_brain1.nii.gz and Test_brain2.nii.gz).
2. Calculate image SNR-related adaptive threshold parameter xp based on the mean and standarddeviation of the 3D-CISS image voxel intensity distribution.
Note: This step takes place inside the main CSF_volumetry() function.
a. Flattening the input image
After removing the obsolete regions using the calculated bounding box, the brain-extracted 3D-CISS input image is flattened into a vector vect, and only non-zero image intensities are considered. This is performed in case any preprocessing step introduces zero or negative values or if other types than MR image are used for analysis (i.e., computed-tomography image that possesses voxels of < 0 in Hounsfield units).
b. Calculation of non-zero intensity distribution
The length, mean, and standard deviation of the vector vect are calculated and saved into matrix V for recording in case of multiple image analysis. A parameter xp is calculated based on the mean (μvect) and standard deviation (σvect) of the non-zero intensities from vect:
x
p
=
σ
v
e
c
t
µ
v
e
c
t
+
σ
v
e
c
t
The parameter xp provides an adaptive correction for the subsequent depiction of CSF space seed regions, reflecting the properties of the distribution of intensities in the brain-extracted 3D-CISS image, i.e., a surrogate for the assessed image SNR.
c. Adjust the predefined percentile threshold of the brain parenchyma intensities distribution in the input image
The initial percentile value P for separating the CSF space from the brain parenchyma image is equal to 95.5, assuming that the CSF space intensities in the brain-extracted 3D-CISS brain parenchyma image are above the 95th percentile of the image’s cumulative distribution of non-zero voxel intensities. Based on the SNR surrogate xp, the adjustment of the percentile P considers three conditions:
i. For images of SNR > 4 (i.e., low influence of Rician noise; CSF space is well-defined in 3D-CISS image, so the percentile P threshold is shifted to higher values: P = 95.5 + xp).
ii. For images of 4 ≥ SNR > 2 (i.e., existing influence of Rician noise; differentiation of the CSF space is affected by noise, but the input image has great T2 contrast, so the separation threshold is unchanged: P = 95.5).
iii. For images of SNR ≤ 2 (i.e., strong influence of Rician noise; differentiation of CSF space is more difficult due to considerable contribution of noise. Differentiation of seed regions requires including a larger part of the voxel intensity distribution, so the percentile P threshold is shifted to lower values: P = 95.5 - xp).
Note: The correction factor xp accounts for subtle intensity changes and does not result in the threshold going below 92nd and exceeding the 97th percentile of the brain-extracted and bias-corrected 3D-CISS image voxel intensity distribution.
3. Perform initial CSF space segmentation by means of an adaptive intensity threshold and calculation of the cumulative distribution of voxel intensities > 0 from the brain-extracted and bias-corrected 3D-CISS volume.
Function Definition and Description
The function [mask] = intensify_CSF(image, P) processes the brain-extracted and bias-corrected 3D-CISS image to adaptively enhance (CSF) space regions and to depict the CSF space seed regions for subsequent automatic region growing segmentation. The function uses intensity distribution normalization and P threshold parameter from the prior step.
Inputs: image (3D image data) and P (percentile value used for adaptive thresholding of the image intensities).
Output: mask (binary 3D mask image highlighting the CSF seed regions for further region growing algorithm).
a. Flattening the input image:
The brain-extracted and bias-corrected 3D-CISS input image is flattened into the vector vect, and the mean and standard deviation of non-zero voxel intensity distribution is calculated as in step 2a (see above). This calculation is repeated in case the user wishes to use the intensify_CSF() function independently.
b. Image intensity distribution normalization:
Each voxel of the input image is rescaled to enhance the distinction of the CSF space regions, assuming that CSF intensities reflect those > 90th percentile of the aggregated non-zero intensities distribution within the image. The image is normalized by subtracting 1.33 times the standard deviation of non-zero intensities (σvect) and then dividing by the σvect. This rescaling factor is applied to ensure that all high-intensity CSF space regions are included in further consideration, along with their closely neighboring regions affected by the partial volume, but not the rest of the brain parenchyma image:
i
n
t
e
n
s
i
f
i
e
d
_
C
S
F
_
i
m
a
g
e
=
i
m
a
g
e
-
1.33
×
σ
v
e
c
t
σ
v
e
c
t
The rescaled (normalized) voxel intensities are saved using a floating point precision in the range between − 1.33 × σvect and the new distribution peak close to the image SNR defined as µvect/σvect (the maximum rescaled intensity ~10 and the mean value varying between 3 and 5 among all analyzed images in the original study [34]).
c. Adaptive thresholding:
All the rescaled voxels possessing negative intensity (i.e., belonging to the brain parenchyma image) are assigned to 0, and a new aggregated distribution of the rescaled voxels of > 0 intensity is computed. Subsequently, the image intensity at the P percentile (see step 2c) of the new non-zero voxel intensity distribution is calculated and denoted as Px. All voxels of intensities ≤ Px are assigned to 0 to keep only the high-intensity CSF seed regions (Figure 1D).
d. Binary CSF space mask conversion:
All remaining non-zero intensities are set to 1, creating a binary mask image where the CSF seed regions are highlighted.
Note: An optional 3D median filtration is available after the intensify_CSF() function is executed (Figure 1E) and is recommended if the computation of mask image leads to the depiction of multiple non-continuous and spurious segments due to low SNR or high influence of Rician noise in the input image. However, it is also recommended to compare the results of segmentation with and without 3D median filtration.
4. Final segmentation: Apply region growing algorithm to reconsider borders of the seed CSF space regions.
Functions Definition and Description
The function [mask2] = grow_regions(mask, image, plt, xp) reconsiders the borders of the CSF space seed regions in the sagittal plane, based on their intensity and contrast to the neighboring regions.
Similarly, subsequently called two consecutive times function [mask3] = grow_regions2(mask2, image, plt, xp, mfilt) reconsiders the CSF space regions in axial and coronal planes, respectively. The function updates the CSF space mask based on the output from the previously called function.
Inputs: mask (binary mask image with initial or updated seed CSF space regions), xp (percentile adjustment for the adaptive intensity thresholding), image (original brain-extracted and bias-corrected 3D image), mfilt (flag for optional slice-wise 2D median filtration), and plt (flag for plotting intermediate results).
Output: mask (binary 3D mask image with reconsidered CSF space image regions).
a. Flattening the input image
The brain-extracted and bias-corrected 3D-CISS input image is flattened into a vector vect, and the mean and standard deviation of non-zero voxel intensity distribution is calculated as in step 2a (see above). As before, this calculation is repeated in case the user wishes to use this function independently from the whole algorithm.
b. Region growing
The function processes each corresponding slice of the image and mask volumes parallelly, excluding the last border slice. For each slice of the image, its original and its zero-padded (extended by n zero-value voxels in both sides of two orthogonal directions, to account for the largest region growing kernel specified below) copies are made to avoid manipulations in the original image. If the analyzed image slice contains any positive values (i.e., CSF regions are present), boundaries of the seed CSF regions are computed in the corresponding slice of the mask volume (Figure 1F–H).
Dilation: To ensure that only voxels belonging to the CSF space and not affected by the partial volume from the surrounding parenchyma are considered, these are included in the final CSF space mask if their intensity values are ≥ (97.5th - xp) percentile (i.e., μd + 2 × σd) of the aggregated voxel intensity distribution from the image and fulfill the contrast calculation-based condition for continuity of the CSF space. The boundary contrast is calculated as an absolute relative CSF to brain parenchyma contrast in the image, considering the mean of n voxels intensities at each side from the boundary voxel in horizontal/vertical/diagonal directions. The contrast is calculated for n from 1 to 4 in the sagittal and from 1 to 3 voxels in the axial and coronal planes. The voxels at nth distance from the boundary are included in the updated CSF mask if their absolute relative contrast is below the threshold α = 2%.
Erosion: Similarly, the updated CSF mask is recalculated, and the voxels are reconsidered for exclusion using the same method. Herein, the contrasts are calculated for a smaller n from 1 to 2 in the sagittal and from 1 to 3 in the axial and coronal planes, to avoid removing small regions belonging to the perivascular space around the main cerebral arteries. The voxels at nth distance are removed from the updated CSF mask if their intensity in the original CISS image is ≤ (95.5th - xp) percentile [for sagittal, and < (95.5th - xp) for the axial and coronal planes] of the aggregated intensity distribution, and the absolute relative contrast to the respective boundary voxel is above α2 = 2.5% (for sagittal, or above and equal to α2 for the remaining planes).
The refined slices are added to the final output mask.
Note: The version of the algorithm equipped with the user interface has additional input structure params allowing for changing the 97.5th percentile threshold as well as the α and α2 parameters so that the user can define the conditions for the region growing algorithm.
c. Optional morphological filtering
If median filtering mfilt is enabled, a 2D median filtration with a 3 × 3 voxels kernel is applied to every slice of the updated CSF space mask image, to remove remaining false-positively segmented single voxels and to enclose wrongly opened larger regions (grow_regions2() function only).
d. Plotting
If plotting plt is enabled, intermediate results are displayed to visualize the CSF space mask refinement process (Figure 1F–H).
5. Saving of the output
The final median-filtered and unfiltered output CSF space mask volumes are saved in NIfTI format into the directory of the original input file. Names of the output files reflect the original input file name appended with “CSF_mask_medfilt_final.nii” and “CSF_mask_final.nii” in case of median-filtered and unfiltered 3D volumes, respectively. After the automatic algorithm successfully finished processing, a confirmation dialog opens in a separate window (Figure 1I).
Note: Execution time for the algorithm depends on the number of processor cores, available memory, image resolution, and the size and number of segmented ROIs. For high-resolution 3D-CISS whole-brain images with a voxel size of (33 μm)3 (as analyzed in [34], image sample provided in the Supplementary information), the processing time in MATLAB ranged from 3.5 to 4.5 minutes using a personal computer equipped with an Intel Core i7-10700U 6-core processor and 32 GB of RAM.
Data analysis
The presented algorithm was intentionally developed for automatic CSF space delineation in high-resolution 3D-CISS images. However, we encourage applying the algorithm for the analysis of other types of MRI or micro-computed tomography images, by interactively changing the input parameters to depict different parts of the image voxel intensity distribution (see user interface in Figure 1A and B). Usage of the algorithm does not require substantial skills in MATLAB; however, any modifications require an understanding of the MATLAB code, organization of the 3D image data matrix, and mathematical reasoning behind the algorithm.
Validation of protocol
The capabilities of the algorithm were tested in 3D-CISS images acquired with (50 μm)3 and (33 μm)3 isometric voxel size, presented during the ESMRMB 2021 conference [40] in the original research paper [34].
General notes and troubleshooting
Usage of 3D-CISS for CSF space imaging
When acquiring 3D-CISS images, it is important to separately consider the acquisition of 3D-TrueFISP volumes, which are significantly affected by banding artifacts. Despite compensatory measures such as increasing resolution, bandwidth per voxel, and reducing both time-to-repetition and time-to-echo, these artifacts remain challenging to eliminate in clinical (low-field) and especially preclinical (high-field) MR systems. They are particularly prominent at the skull base and other regions where magnetic field susceptibility distortions are unavoidable, resulting in an overall loss of image quality. To better address these issues, we recommend acquiring 3D-CISS using four (instead of two) TrueFISP acquisitions, each in orthogonal phase encoding directions (0°, 180°, 90°, and 270°). It is also important to note that our study's evaluation may be affected by the parabolic CryoProbe sensitivity profile. To mitigate this, we recommend performing a bias field correction of 3D-CISS images as we did in the original study [34,40].
Delineation of CSF space in 3D-CISS images using the proposed method
We have demonstrated that our automatic CSF segmentation in 3D-CISS images is effective in depicting subtle differences in CSF volume between wild type and animals lacking the aquaporin-4 channel [34]. With the highest possible spatial resolution achieved using our setup and only two phase-encoding directions applied for CISS image calculation, our results showed good differentiation of the CSF space without the need for contrast agent injection. It is worth noting that using four encoding directions in our setup would increase acquisition time to up to three hours and might introduce additional biases from variability in anesthesia levels or due to prolonged animal restraint. Additionally, single voxel layers, fuzzy or smaller than 33 μm in any dimension regions, might have been missed due to partial volume effects or residuals from banding artifacts. Therefore, besides acquiring TrueFISP in more encoding directions, we also recommend applying more averages (repetitions) for CISS image formation if the protocol time allows. Finally, we encourage researchers to use this algorithm and further validate its action in their images, especially in the presence of a reliable and known segmentation ground truth.
Supplementary information
The following supporting information can be downloaded here:
The algorithm implementation in MATLAB can be downloaded in https://github.com/RSG1UCPH/CSF-space-segmentation-in-3D-CISS.git). Additionally, exemplary 3D-CISS volumes can be found here:
1. “Test_brain1.nii.gz” – high-resolution 3D-CISS volume acquired using (33 μm)3 using voxels volume
2. “Test_brain2.nii.gz” – 3D-CISS volume acquired using (100 μm)3 using voxels volume
3. Figure S1. 3D-CISS volume
4. Figure S2. 3D-CISS image preprocessing steps
Acknowledgments
The algorithm described in the current protocol was originally developed and employed for objective assessment of the volumes and structural differences between the brain CSF spaces in aquaporin-4-null and wild-type littermate mice, based on 3D-CISS images [34]. Development of this algorithm was supported by Lundbeck Foundation (R359-2021-165), Novo Nordisk Foundation (NNF20OC0066419), and the Dr. Miriam and Sheldon G. Adelson Medical Research Foundation.
Competing interests
The authors declare no competing interests.
Ethical considerations
The original research ([34]) that this computational method was developed for received approval from the University of Copenhagen Animal Experiment Inspectorate and the University of Rochester Medical Center Committee on Animal Resources. Development and usage of this method do not require any special approvals. The method does not analyze any details of the subjects that underwent imaging and considers solely a 3D matrix of an original image as an input.
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Article Information
Publication history
Received: Aug 2, 2024
Accepted: Oct 31, 2024
Available online: Nov 14, 2024
Published: Jan 5, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Neuroscience > Basic technology > MRI
Biophysics > NMR spectroscopy > NMR imgaing
Computational Biology and Bioinformatics
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Microfluidic Cultures of Basal Forebrain Cholinergic Neurons for Assessing Retrograde Cell Death by Live Imaging
SD Srestha Dasgupta
MP Mansi A. Pandya
WF Wilma J. Friedman
Published: Vol 15, Iss 1, Jan 5, 2025
DOI: 10.21769/BioProtoc.5149 Views: 244
Reviewed by: Domenico Azarnia TehranAlessandro DidonnaMiaomiao Tian
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Original Research Article:
The authors used this protocol in eNeuro Aug 2023
Abstract
Neurons are highly polarized cells, with axons that may innervate distant target regions. In the brain, basal forebrain cholinergic neurons (BFCNs) possess extensive axons that project to several target regions such as the cortex, hippocampus, and amygdala, and may be exposed to a specific microenvironment in their axon targets that may have retrograde effects on neuronal health. Interestingly, BFCNs express the pan-neurotrophin receptor p75NTR throughout life while also concomitantly co-expressing all Trk receptors, making them capable of responding to both mature and precursor neurotrophins to promote survival or apoptosis, respectively. Levels of these trophic factors may be modulated in the BFCN axon or soma microenvironment under neurodegenerative conditions such as seizure and brain injury. In this protocol, BFCNs are established in microfluidic devices for compartmental culture, with the aim of studying the effects of axon- or soma-specific stimulation of BFCNs for an in vitro representation of distal axon vs. soma environments as seen in vivo. This study further establishes a novel method of tracing and imaging live BFCNs exposed to stimuli in their distal axons with the aim of assessing retrograde cell death. The in vitro compartmental culture system of BFCNs that allows live imaging may be applied to investigate various effects of axon- or soma-specific stimuli that affect BFCN health, maintenance, and death, to model events that occur in the context of brain injury and neurodegenerative disorders.
Key features
• Separation of axons and soma of basal forebrain primary neurons in vitro using microfluidic chambers.
• Compartmental/localized treatment of axons or somas of BFCNs.
• Live imaging of retrogradely labeled BFCNs to assess cell death.
Keywords: Microfluidics Basal forebrain Live imaging Cell death
Background
Neurons may possess long axons that project to distal areas and respond retrogradely to stimuli limited to their axonal microenvironment. Basal forebrain cholinergic neurons (BFCNs) send extensive and long axonal projections to several distal brain areas, such as the cortex, to regulate cognitive functions such as attention, emotion, and memory [1]. Interestingly, throughout their life, BFCNs express the pan-neurotrophin receptor p75NTR as well as all three members of the Trk family of receptor tyrosine kinases. Neurotrophins are essential for BFCNs’ survival, maintenance, differentiation, and function [2]. Neurotrophins in their precursor and mature forms activate different signaling cascades, which lead to contrasting cellular responses. Proneurotrophins are cleaved to form mature neurotrophins, which promote a pro-survival response through Trk signaling [3–6]. In contrast, under conditions of brain insult such as seizures or traumatic brain injuries (TBI), proneurotrophins (proNTs) bind p75NTR and sortilin to promote apoptosis in BFCNs [7]. Interestingly, as BFCNs express all the neurotrophin receptors, these neurons are capable of responding to a changing balance of pro vs. mature neurotrophins. To add to the complexity, the dynamic shift in pro vs. mature neurotrophin balance may occur at the BFCN neuronal target regions such as after cortical injury [8]. Although it has been established that p75NTR promotes BFCN degeneration in mass cultures and after seizure conditions in vivo [7], given the loss of BFCNs after cortical injury it is important to answer whether p75NTR can promote BFCN degeneration retrogradely as well, especially when the BFCN soma is unexposed to induced proneurotrophins [8].
This protocol has been applied by Dasgupta et al. [8] to establish whether the changes in the neurotrophic environment of BFCN axon terminals after TBI have a retrograde effect on survival of these afferent projections by live imaging and cell death analysis of in vitro compartmentally cultured BFCNs. Silicone microfluidic devices allow for the segregation of neuronal axons and soma as well as for compartmental manipulation with localized treatments [9], having been used extensively on several neuronal cell types to investigate a myriad of cell biology questions [10]. Moreover, the transparent material allows for labeling and imaging of live tracers that indicate dying cells after treatments. Major studies have established retrograde degeneration through p75NTR in peripheral neurons [11,12], but this protocol helped establish the role of p75NTR and proneurotrophins on BFCN retrograde degeneration, adding to the evidence that suggests that brain injury induced elevation in cortical proneurotrophin leads to p75NTR signaling in BFCNs initiated at the axon terminals to promote retrograde degeneration [8]. Overall, this protocol can be applied to study cell death and other cellular responses to compartmental stimulation of BFCNs by live imaging and can also be adapted to other neuronal populations.
Materials and reagents
Biological materials
1. C57BL/6, strain# 000664 mice were purchased from The Jackson Laboratory and mated in-house. Timed pregnant mice at a pregnancy age of E15 were used for the basal forebrain dissection.
Reagents
1. Poly-d-lysine (Sigma-Aldrich, catalog number: A-003-M)
2. Glucose (Sigma-Aldrich, catalog number: G7021)
3. Transferrin (Sigma-Aldrich, catalog number: T4382)
4. Insulin (Sigma-Aldrich, catalog number: I5500)
5. Putrescine (Sigma-Aldrich, catalog number: P7505)
6. Selenium (Sigma-Aldrich, catalog number: S-5261)
7. Progesterone (Sigma-Aldrich, catalog number: P0130)
8. Penicillin and Streptomycin (10,000 IU) (Sigma-Aldrich, catalog number: P4333)
9. Minimum essential medium (MEM) (Gibco, catalog number: 11095080)
10. Ham’s F-12 media (Gibco, catalog number: 11765054)
11. B-27TM Plus supplement (50×) (Gibco, catalog number: A3582801)
12. Sterile PBS (1×) (Gibco, catalog number: 10010023)
13. Cholera toxin subunit B (recombinant), Alexa FluorTM 488 conjugate (CTB) (Invitrogen, catalog number: C34775)
14. Propidium iodide (PI) (Molecular Probes, catalog number: P1304MP)
15. Sylgard 184 Silicone Elastomer kit (Dow, catalog number: 4019862)
16. Isopropanol (Sigma-Aldrich, catalog number: I9030)
17. Paraformaldehyde (Sigma-Aldrich, catalog number: P6148)
Solutions
1. Serum-free media (SFM) (see Recipes)
Recipes
1. Serum-free media (SFM)
1:1 mixture of Eagle's MEM and Ham's F-12 supplemented with glucose (6 mg/mL), progesterone (20 nm), putrescine (60 μm), transferrin (100 μg/mL), selenium (30 nm), penicillin (0.5 U/mL), and streptomycin (0.5 μg/mL) [4]. Alternatively, you may use commercially available Neurobasal media.
Laboratory supplies
1. Sterile Petri plates (60 mm) (NuncTM EasYDishTM Dishes, catalog number: 150462)
2. Sterile surgical tweezer set (DUMONT Fine Science Tools, catalog number: 11251-10)
3. Surgical scissors (Millipore Sigma, catalog number: S3146)
4. Surgical scalpels (WPI Scalpel Handle, model: #4, catalog number: 500237-G)
5. Sterile scalpel blades (Cincinnati Surgical, catalog number: 00SME11)
6. Spatula (Sigma-Aldrich, catalog number: Z513342)
7. Rapid core puncher, ID 6.0 mm, OD 6.5 mm, white (Well Tech, Ted Pella Inc, catalog number: 15115-12)
8. Hemocytometer (Fisher Scientific, catalog number: 02-671-51B)
9. Sterile serological pipettes (VWR, catalog number: 89130-898, 10 mL)
10. Sterile glass coverslips (25 mm) (Deckglaser, Carolina coverglass, catalog number: 633037)
11. Falcon tube (15 mL) (Fisher Scientific, catalog number: 14-959-53A)
Equipment
1. Laminar flow culture hood (Edgegard) (GMI, catalog number: 8038-30-1041)
2. Nikon SMZ1000 dissecting microscope (Nikon, catalog number: SMZ1000-P)
3. CO2 incubator (Sanyo, model: MCO-17AC)
4. Isotemp oven (Fisher Scientific, catalog number: 13-247-750G)
5. Vertical rotating mixer (Atlantis Bioscience, catalog number: VM-80)
6. Scienceware® vacuum desiccator (Sigma-Aldrich, catalog number: Z119016)
7. Confocal microscope for live imaging (Zeiss, model: LSM 510 microscope)
Software and datasets
1. Fiji: ImageJ (Open Source; https://github.com/imagej) (https://imagej.net/software/fiji/)
Procedure
A. Basal forebrain primary neuron compartmental culture
1. Microfluidic device casting
a. Preparation of positive relief microfluidic mold
i. To create microfluidic devices in-house, Polydimethylsiloxane (PDMS), a silicone elastomer (Sylgard 184), is cast on a positive relief resin microfluidics mold with multiple chambers etched (adapted from Harris et al. [13]; Taylor et al. [9]).
ii. Molds can be reused to cast devices, and therefore need deep cleaning before every casting process. To clean the mold, release pressurized air on the positive relief surface in 5–10 repeats vigorously to remove the remaining PDMS from earlier usage. A thin tube is attached to the pressurized air inlet to which a 1,000 mL pipette tip is attached, and the pipette tip is moved over the surface to create sharp blasts of air; the purpose is to remove any debris from the mold.
iii. Measure the mold volume by adding DI water into the mold area to fill it completely; then, remove the water into a 50 mL Falcon tube. Retain this information for the PDMS preparation and casting step.
iv. Fill the mold area with isopropanol. Put the mold on a shaker at medium speed for 20 min for the isopropanol wash.
v. Remove isopropanol and dry the mold uncovered in an oven at 70 °C for 10 min or until completely dry.
vi. Cover the mold to avoid any particulates adhering to the mold.
b. PDMS preparation and casting
i. Sylgard 184 comprises a PDMS base and a PDMS curing agent, which, when combined, produce the silicone elastomer.
ii. Combine PDMS base and PDMS curing agent in a 50 mL Falcon tube in a 10:1 ratio. Use the precalculated mold volume as a reference to the total volume of the PDMS mixture to be prepared.
iii. Place the mixture on a rotator at low speed for 20 min or until the mixture becomes homogenous.
iv. Pour the PDMS mixture into the clean molds, start at a corner, and let the mixture flow across the surface to fill the entire mold.
v. To remove air from the mixture, place the mold with the mixture in a vacuum desiccator at full suction power for 4–5 h.
vi. Assess the progress of the desiccation by observing the number of bubbles visible in the mixture in the mold every 2 h.
vii. Once the PDMS appears clear and has no bubbles left, gently disassemble the desiccator to ensure the PDMS mixture does not spill out of the mold, as it will still be in a liquid state.
viii. Remove the mold with PDMS and transfer it to an oven at 70 °C for baking. Bake the PDMS for 48 h and keep the mold uncovered.
c. Cleaning and preparation of casted PDMS
i. After 48 h of baking, remove the mold from the oven, cover the mold, and let it cool to room temperature.
ii. Use a spatula with a flat tapered end and a sharp scalpel to extract the solid PDMS from the mold. Start by releasing the edges by cutting with the scalpel, then release the PDMS surface facing the positive relief of the mold with the spatula, and gently peel off the entire PDMS.
iii. Based on the mold structure, each PDMS casting produces nine microfluidic chambers. At this stage, all the chambers are present on the PDMS as one block and need to be separated into individual devices.
iv. Use reusable biopsy punchers (size No. 6) as shown by dotted lines in Figure 1a–c to punch out four holes on the edges of the microfluidic channels of each chamber connected by the microgroove, two on either side, just covering the edges so that an open flow of media will be maintained in both channels.
v. Carefully, cut out individual chambers using the scalpel. Cut the corners of the devices so that the chambers fit 25 mm coverslips optimally and have no overhangs.
vi. Clean out loose PDMS in the chambers using scotch tape on the negatively etched PDMS surface.
vii. Further clean the chambers by washing with 70% ethanol for 10 min.
viii. Dry the chambers at 70 °C for 10 min and store in a sterile Petri dish.
Figure 1. Preparation of microfluidic device by PDMS casting. (a) Schematic of a microfluidic chamber with microgrooves connecting the two compartments. Dotted lines represent the positions to punch holes into the chamber. (b) A microfluidic device made in-house by PDMS casting and preparation using (c) a well puncher of 6 mm inner diameter. Scale bar = 5 mm.
2. Microfluidic chamber preparation for cell culture
a. Microfluidic devices created in-house have two channels connected by microgrooves of 450 μm length, with two compartments on either end of the channels that work as media reservoirs.
b. Perform all further steps in the laminar flow hood to maintain sterility.
c. Prepare individual microfluidic devices a day before cell culture by sterilization with 70% ethanol.
d. Prepare SFM a day before culture (see Recipes).
e. Place individual glass coverslips in 35 mm Petri dishes. Precoat glass coverslips for culture a day in advance with 1 mL of poly-d-lysine (0.2 mg/mL) and maintain overnight at a 37 °C, 5% CO2 cell culture incubator.
f. On the day of the culture, remove poly-d-lysine from coverslips by suctioning carefully, aiming to remove all the solution but not scratch the coated surface.
g. Ensure sterilized microfluidic devices are completely dry as well. Place the negatively etched surface of the microfluidic device on the coverslip with forceps, while pressing gently at each corner to ensure a water-tight seal.
3. Basal forebrain culture
a. Collection of E15 mouse embryo
i. Euthanize a pregnant mouse while the fetuses are at embryonic day 15 (E15) by exposure to CO2.
ii. Sterilize by soaking in 70% ethanol for 5 min.
iii. To remove fetuses under sterile conditions by C-section, make a midline incision in the abdomen to sever the skin using a thick scissor from the surgical tool kit. Sever the muscle along the same incision to expose the embryos.
iv. Remove the embryos using a small scissor and forceps and place in a 20 cm sterile Petri dish with PBS. Maintain the Petri dish on ice.
b. Basal forebrain dissection
i. To collect the basal forebrains, place embryos in SFM to dissect as described below (adapted from Friedman et al. [4]) in the laminar flow hood.
ii. Remove each embryo from its amniotic sac with two straight-edged fine forceps. Separate the heads with the forceps, place in another Petri dish with PBS, and place on ice.
iii. Place a Petri dish with PBS under the dissecting microscope. Place a separated head upright using another set of fine forceps (Figure 2a). Anchor the head on the ventral side with one forceps while clasping onto the top of the skull (gelatinous and soft at this age) with the other forceps. Let go of the ventral anchor and clasp an adjacent section of the skull. Gently peel off the skull with the two forceps with a pull in opposite directions.
Figure 2. Step-by-step basal forebrain dissection captured using the Nikon SMZ1000 dissecting microscope. (a, b) An E15 mouse brain with and without the skull. (c) Mouse brain with the ventral side up, with rostral to caudal ends facing left to right. (d) Demarcations (red) for coronal incision to remove olfactory bulbs and anterior poles. (e) Demarcations for coronal incision at rostral edge of hypothalamus. (f) Red arrow indicates turning the plate 90° clockwise so that the rostral aspect faces anterior. (g) Red curved arrow indicates laying down the coronal slab at 90° toward the posterior direction such that the rostral aspect will face upward. (h) Coronal slab showing the basal forebrain (red circles). (i) Demarcations (red) for dorsal incision at lateral ventricles of both hemispheres. (j) Demarcations (red) for ventral incision to separate the basal forebrain and medial septum. (k, l) Separated basal forebrains of both hemispheres. Scale bar = 1 mm.
iv. Successfully removing the skull will reveal the brain (the shiny translucent layer covering the brain will have been removed) (Figure 2b). Gently turn the head to its side and slide out the brain. Cut the connections at the olfactory bulbs to free the brain from the head with a pair of sharp scalpels.
v. Align the brain with the ventral aspect facing upward and the olfactory bulbs on the rostral aspect of the brain to the left (Figure 2c).
vi. Cut coronally to remove the frontal poles of the cortex along with the olfactory bulbs (Figure 2d).
vii. Cut at the demarcation of the rostral edge of the hypothalamus to generate a thick coronal section (Figure 2e).
viii. Turn the thick coronal section at 90°, laying the section down such that the rostral cut side is facing upward (Figure 2f, g).
ix. In the coronal section, make incisions at the dorsal tips of the lateral ventricle on both hemispheres (Figure 2h, i) and at the ventral surface of the brain to separate the medial septum, together with portions of the diagonal band of the basal forebrain (Figure 2j). Collect the separated septum and diagonal band of the basal forebrain (Figure 2k, l) and clear the rest of the tissue debris to one side of the Petri dish.
x. Repeat steps v–ix to collect the basal forebrain from all the embryos into a separate small Petri dish (30 mm) with SFM.
xi. Cut the collected tissue into small pieces with the scalpels in a low volume of media in a separate Petri dish (500 µL to 1 mL), aiming to have a high concentration of cells (approximately 2–3 million cells/mL). Dissociate cells using a Pasteur pipette by trituration (repeated pipetting) approximately 20–30 times until cells are dissociated into a cloudy suspension.
c. Plating BFCNs in microfluidic devices
i. Count cells using a hemocytometer and pipette 50,000 cells in a pressure pulse through one side of the microfluidic chamber that should populate the channel. This channel will be considered the soma compartment. The average cell yield per basal forebrain (both hemispheres combined) per embryo is about 800,000 cells.
ii. Basal forebrains pooled from several embryos dissociated in 500 μL–1 mL of media should yield an optimal concentration of cells (2–3 million/mL) to plate 50,000 cells in a concentrated media volume (~10 μL).
iii. Prepare SFM + 1% B27 to plate and maintain cells.
iv. After 5 min of plating cells, add 200 μL of SFM + 1% B27 media to each reservoir of the soma compartment. Add 100 μL of SFM + 1% B27 in the reservoirs of the opposite side, which is considered the axon compartment.
v. Maintain a volume gradient between the soma and axon side to promote neurite growth through the microgrooves to the axon compartment for five days at 37 °C and CO2 (5%) to obtain compartmentalized BFCN cultures.
vi. Add 50 μL of SFM + 1% B27 to all compartments every alternate day after culture to account for media evaporation from the microfluidic devices.
B. In vitro labeling and live imaging of BFCNs in microfluidics
1. Retrograde labeling of live neurons
a. Prepare a working solution of a retrograde tracer, Alexa 488 labeled choleratoxin B (CTB, 1 μg/mL) in SFM.
b. At 5 days in vitro (DIV), treat the axon compartment with the SFM + CTB working solution and incubate for 20 min in the incubator.
c. After 20 min, wash the axon compartment twice with SFM + 1% B27 media to remove any excess tracer, then add 100 μL of media to maintain axons.
d. Incubate for at least 5 h to allow CTB to be transported retrogradely to the somas of BFCNs that have extended their axons into the distal compartment.
e. Prior to compartmental stimulation of the BFCNs with desired treatments, replace the soma media with a working solution of SFM + 1% B27 + PI (1 μg/mL) for 10 min. PI will label the nuclei of dying neurons with an excitation maximum of 535 nm. PI does not need to be washed out and remains in the media throughout the experiment.
f. Maintain the media volume gradient between the soma and axon compartment for the duration of the treatment as well.
2. Live imaging of microfluidics to assess cell death
a. For live imaging, cells need to be maintained at a constant temperature (37 °C) and CO2 (5%) for the duration of imaging for the experiment.
b. Set up the microscope equipped for live imaging with these settings before adding treatments to the cell culture.
c. Set up the microscope at 10× objective. Use the brightfield channel to locate the microgrooves such that both the axon and soma compartments are in the optical field (Figure 3a).
Figure 3. Live tracing and imaging of basal forebrain cholinergic neurons (BFCNs) in microfluidic cultures to study retrograde cell death. (a) Representative images of E15 mouse basal forebrain neurons cultured in microfluidic chambers for 5 DIV labeled with a retrograde tracer CTB Alexa 488. Before live imaging, propidium iodide (PI) was added to the soma compartment to identify dying neurons. Scale bar = 50 μm. (d) After 24 h of treatment, CTB+ (white arrowheads) cells are counted as surviving cells, whereas CTB+PI+ (blue arrowheads) cells are counted as dying cells. Scale bar = 20 μm.
d. CTB 488 has an Alexa 488 fluorophore attached to it, and PI fluoresces at a 555 nm wavelength once it enters the nuclei of dying cells. Set up the 488 and 555 channels at a low laser power (5%–10%) to avoid phototoxicity optimized based on signal from the sample. Adjust digital offset and gain if necessary, using the range indicator. Set camera settings to capture images at 1,024 × 1,024 pixel size, at a scan speed of 8. Maintain these settings for the entire experiment. Optimization is required for each biological replicate.
e. Right after adding treatments, capture fluorescence images of the soma compartment of the microfluidics with excitation at 488 nm and 555 nm to capture CTB and PI, respectively, at 0 h.
f. Return the cells to the incubator for 24 h. Alternatively, for continuous live imaging, maintain the cells in the live imaging chamber on the microscope and capture images at different time points appropriate for the experiment.
g. Image the live cells in the soma compartment as mentioned above at the 24 h time point.
h. Once live imaging experiments are completed, fix cells with 4% paraformaldehyde solution while keeping the microfluidic device attached to the coverslip, if further immunolabeling post-fixation is intended. Then, the microfluidic device can be removed for further processing of the coverslips.
Data analysis
This experimental protocol can be applied to understand the effects of axon-specific stimulation with ligands of choice on cell death, as PI does not incorporate into live cells. Therefore, we may assess the effect of compartmental treatments on neurons that are labeled with the retrograde tracer CTB but not PI at 0 h, by analyzing PI incorporation in these neurons at 24 h. CTB-positive neurons that do not incorporate PI in their nucleus at 24 h are considered surviving neurons, whereas CTB and PI double-positive neurons are considered dying neurons in response to the treatment (Figure 3a, b).
The retrograde degenerative effect of axonally sourced proneurotrophins in BFCNs was reported by Dasgupta et al. [8] using this model system and live imaging analysis. In this study, axonal treatment of BFCN microfluidic cultures with the proneurotrophins proNGF and proBDNF resulted in retrograde degeneration, in comparison to untreated control cells [8]. Live images of the entire soma compartment, CTB, and PI channels, were captured at 10× in an LSM 510 confocal microscope at 0 and 24 h. Microfluidic chambers with a minimum of 50 CTB-positive cells were considered for the analysis. Each treatment in every biological replicate had two technical replicates averaged for the final analysis. Quantification was done in ImageJ by counting PI in CTB+ve cells, which was normalized as a percentage of PI+CTB+ cells over CTB+ve cells [8]. A higher percentage indicates an increase in cell death.
Validation of protocol
This protocol was used to establish the neurodegenerative effects of axon-specific stimulation of basal forebrain neurons with proneurotrophins by Dasgupta et al. [8]. In this study, it was shown that cortical injury promotes a retrograde degeneration of basal forebrain neurons through p75NTR. Moreover, it was established that cortical injury leads to an induction of proneurotrophins in the cortex but not in the basal forebrain, suggesting that p75NTR binds proneurotrophins at the BFCN axon terminals that innervate the cortex, which leads to retrograde degeneration. In vitro live imaging of microfluidic BFCN cultures using the described protocol showed that axon-specific proneurotrophin stimulation for 24 h promoted cell death [8], further supporting the hypothesis that axonally sourced proneurotrophins are sufficient to promote retrograde degeneration of BFCNs.
General notes and troubleshooting
A. Microfluidic chamber preparation
1. Instead of making in-house devices, commercial silicone microfluidic devices may be purchased from Xona Microfluidics (XC450), which can be used with the same method.
2. The PDMS mixture should completely fill the mold during chamber preparation, as less PDMS will lead to the formation of a device with less thickness, making it too malleable and prone to breaking even though the negative relief of the microfluidic device will be functional.
3. The percentage of PDMS base to curing agent may be changed to produce a device with more or less elasticity. However, the proportion used was optimized in-house to have the most effective devices, which have adequate elasticity to form a complete seal with the glass coverslip, while not being too elastic, which hinders releasing the coverslips from the chambers for postprocessing such as immunostaining.
4. Post-rotating the PDMS mixture will have a lot of bubbles. Though the long desiccation process should help remove all the bubbles over time, spinning down the mixture in a centrifuge may aid in removing bubbles more effectively before the PDMS is poured into the molds, as well as shorten the duration of the desiccation step.
5. While creating media reservoirs in freshly prepared microfluidic devices, place punchers as close as possible to the edges of the compartments to ensure an open channel for media between the compartments. Using a sharp scalpel, the edge leading into the compartment can be shaved slightly to increase the opening, which reduces the fluid pressure and allows smooth media flow.
6. To maintain a water-tight seal between the microfluidic chamber and the coverslip it is placed on, both need to be dry at the time of assembly, as any moisture on either leads to a break in the seal, which hinders axon–soma separation.
B. Basal forebrain dissection and live imaging
1. It is important to have a minimum concentration of cells of 2–3 million/mL in the plate microfluidics. The total number of cells plated (50,000 cells) should be optimally present in a volume of 15 μL. A minimum of 8 μL to a maximum of 30 μL media volume is advisable. Below the minimum volume, cells may not optimally pass through the soma compartment and be concentrated on one side. Above the maximum volume, the cell density is too dilute to populate the soma compartment and may flow through to the other side. The aim is to condense the cell plating in the soma compartment and not the reservoirs.
2. In case of low cell concentration, collect cells in a sterile 15 mL Falcon tube and add 8 mL of media (SFM + 1% B27) to the cells. Centrifuge the cells at 400× g for 5 min at 4 °C to collect the cells in a pellet. Carefully remove the supernatant and resuspend the cells in an appropriate volume of media to obtain the optimal cell concentration as discussed above. Follow up with plating cells as described in the procedure (step A3c).
3. Methods of enzymatic dissociation such as trypsinization or DNase treatment may be chosen if the quality of the cell culture is being affected by the mechanical dissociation step. However, at this developmental age, the basal forebrain tissue is soft and should ideally not require it.
4. Incubation time after treating with CTB can be extended from 5 h to overnight in case the CTB signal is not prominent after 5 h to allow transport of the CTB cargo to the soma from the axon.
Acknowledgments
Funding sources for this study are NIH/NINDS 1R01NS127894, Busch Biomedical Grant Program, and SASN Innovation and Development Award to WJF. This protocol was originally described and validated in an eNeuro research paper titled “Cortical Brain Injury Causes Retrograde Degeneration of Afferent Basal Forebrain Cholinergic Neurons via the p75NTR” by Dasgupta et al. [8]. Dr. Eran Perlson is thanked by the authors for the molds for the microfluidic chambers.
Competing interests
The authors declare no competing interests.
Ethical considerations
All experiments were approved by Rutgers University, Newark and performed in compliance with the Institutional Animal Care and Use Committee (IACUC) policies.
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Publication history
Received: Aug 12, 2024
Accepted: Oct 30, 2024
Available online: Nov 17, 2024
Published: Jan 5, 2025
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© 2025 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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Using HBmito Crimson to Observe Mitochondrial Cristae Through STED Microscopy
XG Xichuan Ge *
WR Wei Ren *
CS Chunyan Shan
PX Peng Xi
BG Baoxiang Gao
(*contributed equally to this work)
Published: Vol 15, Iss 1, Jan 5, 2025
DOI: 10.21769/BioProtoc.5150 Views: 269
Reviewed by: Domenico Azarnia TehranAftab NadeemRozemarijn Van Der Veen
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Original Research Article:
The authors used this protocol in light: science & applications May 2024
Abstract
Mitochondrial cristae, formed by folding the mitochondrial inner membrane (IM), are essential for cellular energy supply. However, the observation of the IM is challenging due to the limitations in spatiotemporal resolution offered by conventional microscopy and the absence of suitable in vitro probes specifically targeting the IM. Here, we describe a detailed imaging protocol for the mitochondrial inner membrane using the Si-rhodamine dye HBmito Crimson, which has excellent photophysical properties, to label live cells for imaging via stimulated emission depletion (STED) microscopy. This allows for STED imaging over more than 500 frames (approximately one hour), with a spatial resolution of 40 nm, enabling the observation of cristae dynamics during various mitochondrial processes. The protocol includes detailed steps for cell staining, image acquisition, image processing, and resolution analysis. Utilizing the superior resolution of STED microscopy, the structure and complex dynamic changes of cristae can be visualized.
Key features
• The protocol is designed to visualize mitochondrial cristae in living cells using STED microscopy.
• The protocol enables nanoscale observation of dynamic mitochondrial cristae.
• Real-time observation of mitochondrial morphological changes, fusion, and fission events.
Keywords: Live cell imaging Super-resolution imaging Mitochondria cristae Low-saturation power Fluorescence labeling
Graphical overview
Background
Mitochondria are crucial organelles responsible for ATP production, metabolic regulation, calcium homeostasis, and participation in cellular signaling processes [1–5]. The mitochondrial inner membrane (IM) is a crucial structure within mitochondria, containing the oxidative phosphorylation complexes responsible for ATP synthesis. These complexes on the IM play a key role in cellular energy supply by hosting the entire respiratory chain [6,7]. The IM folds into cristae to increase the surface area, thereby supporting greater ATP production. The shape of the cristae is altered by mitochondrial dynamics, which can affect mitochondrial respiration [8].
The diameter of mitochondria ranges from 200 to 700 nm, with cristae spacing typically around 70 nm [9]. Conventional confocal microscopy has a resolution limited to approximately 200 nm, which is insufficient to resolve the details of these structures; however, the advent of super-resolution microscopy has enabled the visualization of cristae [10]. Stimulated emission depletion (STED) microscopy, in particular, achieves high resolution by employing a doughnut-shaped depletion laser to selectively erase peripheral fluorescence following sample excitation, thereby improving the point spread function (PSF). STED technique provides a spatial resolution of 50 nm and temporal resolution of 1 frame per second, making it a powerful tool for dynamic imaging of mitochondrial cristae at the single-crista level.
In the STED system, the intensity of the depletion beam is several thousand times higher than that of the excitation beam, requiring fluorophores in the fluorescent probes to possess exceptional photostability to avoid quenching. Therefore, we have designed a novel fluorescent dye for STED imaging. HBmito Crimson is characterized by exceptional photostability, targeted accumulation on the mitochondrial IM, and emission only upon binding to the IM, as shown in Figure 1A. In pure organic solvents or water, HBmito Crimson exhibited its absorption peak at 660 nm and an emission peak at 688 nm (Figure 1B). The emission spectrum tail of HBmito Crimson extends toward 775 nm, generating a small peak near this wavelength. This reduces the dye’s saturation power, significantly enhancing the depletion efficiency of the 775 nm laser [11].
Figure 1. Structural and spectral properties of HBmito Crimson for mitochondrial labeling. A. The chemical structure of HBmito Crimson is used for the specific labeling of the mitochondrial inner membrane. B. The absorption and emission spectra of HBmito Crimson, which can be depleted using a 775 nm laser.
In this protocol, we describe in detail the preparation of cells, labeling methods with HBmito Crimson, image acquisition, and image processing methods suitable for confocal and STED microscopy imaging.
Materials and reagents
Biological materials
1. COS7 cell (ATCC, CRL-1651)
Reagents
1. Dulbecco’s modified Eagle’s medium (DMEM) (Gibco, catalog number: 11965-092), store at 4 °C
2. Fetal bovine serum (FBS) (Gibco, catalog number: 10091148), store at -20 °C
3. Penicillin/streptomycin solution (Gibco, catalog number: 15140122), store at -20 °C
4. Trypsin-EDTA solution (Gibco, catalog number: 25200056), store at -20 °C
5. Dulbecco’s phosphate buffered saline (PBS) (Gibco, catalog number: 11965-092), store at 4 °C
6. HBmito Crimson, 200 μM DMSO solution, store at -20 °C
The dyes in this article can be synthesized and purified according to the literature [11] or purchased from MedChemExpress (catalog number: HY-D2346).
7. DMSO (analytical reagent grade) (Aladdin, catalog number: D103272), store at room temperature
8. Immersion oil (Olympus, catalog number: IMMOIL-F30CC)
Solutions
1. Complete COS7 cell media (see Recipes)
2. HBmito Crimson solution (see Recipes)
3. HBmito Crimson DMSO solution (see Recipes)
Recipes
1. Complete COS7 cell media
Reagent Final concentration Amount
DMEM n/a 44.5 mL
FBS 10% 5 mL
Penicillin/streptomycin solution 1% 0.5 mL
Total n/a 50 mL
The solution should be stored at 4 °C and used within one month. Ensure that it is sterile each time it is used. FBS should be filtered through a 0.2 μm filter membrane.
2. HBmito Crimson solution
Reagent Final concentration Amount
HBmito Crimson, 200 μM 500 nM 2 μL
COS7 cell media n/a Up to 1 mL
Total n/a 1 mL
Prepare the solution before each use.
3. HBmito Crimson DMSO solution
Reagent Final concentration Amount
HBmito Crimson, 5 mM 5 μM 1 μL
DMSO n/a Up to 1 mL
Total n/a 1 mL
Prepare the solution before each use.
Laboratory supplies
1. Sterile 1.5 mL polypropylene centrifuge tubes (any vendor)
2. Sterile 50 mL polypropylene centrifuge tubes (any vendor)
3. Sterile nuclease-free filter tips (10, 200, and 1000 μL) (any vendor)
4. 35 mm glass bottom dishes, 170 μm ± 5 μm (Standard Imaging, catalog number: STGBD-035-1; Cellvis, catalog number: C35-20-1.5H)
5. Pipette tips, sterile (any vendor)
6. Cell culture flasks, vented flask surface area 25 cm2 (any vendor)
7. Pipettor (Eppendorf Research® plus) with volume ranges of 0.5–10 μL, 10–100 μL and 100–1000 μL
8. 0.2 μm filter membrane (Millex-GP, catalog number: SLGPR33RB)
Equipment
1. Abberior Facility Line (Abberior Instruments GmbH, Germany) with a 60× oil immersion objective (N.A. 1.42, Olympus, Japan)
2. Common lab equipment: cell culture incubator, safety cabinet, -20°C freezer, fridge (any vendor)
3. Microscope incubator (OKOlab, model: H301-T-UNIT-BL-PLUS)
Software and datasets
1. Abberior Imspector (16.3.14280)
2. Huygens SVI (23.10)
3. ImageJ Fiji, version 2.1.0. with Java 1.8.0_172
4. Origin (2019b)
Procedure
A. STED saturation power measurement
STED saturation power refers to the laser power required to effectively deplete the fluorescence of dye molecules. Dyes with low saturation power improve the overall quality of STED imaging, especially in high-resolution, long-term experiments. Therefore, saturation power is a key parameter for determining whether a dye is suitable for high-resolution imaging. Additionally, measuring saturation power can help estimate the optimal depletion laser intensity needed for imaging, such as in section C for setting the depletion power of HBmito Crimson. The resolution in STED microscopy can be described by the following formula:
d
≈
λ
2
N
.
A
.
1
+
I
S
T
E
D
I
s
a
t
λ is the excitation wavelength, N.A. is the numerical aperture of the microscope objective, ISTED is the intensity of the depletion laser, and Isat is the saturation intensity of the fluorophore. From the formula, it is evident that when ISTED remains constant, decreasing the saturation power Isat increases ISTED/Isat, resulting in higher resolution. Therefore, low-saturation-power probes can achieve high resolution at lower laser powers, effectively reducing photobleaching and phototoxicity.
1. Add 1 mL of HBmito Crimson DMSO solution to a 35 mm glass bottom dish.
2. Set the spatial light modulator (775 nm) mode to “none” in the background interface of the commercial STED system, transforming the depletion beam of the donut distribution into a Gaussian distribution. Set the excitation intensity to 3%, which can be calibrated according to the dye's brightness. Adjust the excitation intensity to 3%, calibrated based on the brightness of the dye. Gradually increase the depletion laser intensity from 0%, in increments of 1%–3% (approximately 2–5 mW), e.g., 0%, 2%, 4%, and so on, up to 50%. At each depletion laser intensity level, capture two images—one with the excitation laser on and one with it off. Record the average fluorescence intensity for each image.
3. For each depletion laser power, subtract the average fluorescence intensity of the excitation-off image from that of the excitation-on image. Plot the normalized fluorescence intensity against the depletion laser intensity. Fit the data to a curve and identify the point at which the fluorescence intensity decreases to 50% of its initial value. The corresponding depletion laser intensity at this point is the saturation power of the dye.
Note: Repeat the imaging and recording at each laser intensity multiple times to ensure data accuracy and reliability. Monitor the sample for photobleaching to prevent damage from excessive excitation and adjust the laser settings as needed to minimize photobleaching.
B. Sample preparation
1. In our work, COS7 cells were cultured in a cell culture flask with COS7 medium. The cells were maintained at 37 °C in a humidified atmosphere containing 5% CO2 for 2–3 days.
2. When the cells reach 80% density, remove the original COS7 medium and wash the cells with 3 mL of PBS to eliminate the residual medium. Add 1 mL of trypsin-EDTA solution for digestion for 3 min. After digestion, add 1 mL of COS7 cell medium to stop the digestion process.
3. Centrifuge the digested cells at 200× g for 3 min to collect the cell pellet.
4. Seed approximately 6 × 104–9 × 104 cells into 35 mm glass-bottom dishes. For optimal imaging results, ensure that the cells are evenly distributed and not overcrowded after seeding.
5. Allow the cells to grow in the incubator for 24 h. A confluence between 40%–80% is ideal for subsequent labeling and imaging procedures. If cell washing is required, the optimal imaging time is within 1 h after washing.
6. Remove the original medium and add 1 mL of HBmito Crimson solution, incubating for 10 min in a 37 °C, 5% CO2 environment. Without washing, proceed directly to STED imaging, making the necessary adjustments to meet specific experimental requirements.
Note: When preparing cells for imaging, several key factors must be considered:
Cell density: Maintain an appropriate cell density (between 40% and 80%) to ensure cell health while providing sufficient space for single-cell imaging. High cell density may lead to cell aggregation, adversely affecting image quality.
Culture medium selection: Choose an appropriate basal medium and serum for different cell lines to ensure optimal growth conditions.
Contamination control: Cells must be free of any contamination to ensure reliable imaging results. The inclusion of antibiotics in the culture medium can help prevent contamination.
C. Image acquisition
This protocol is specifically designed for use with the Abberior Facility Line, but the method can be adapted for use with other STED microscopes as well.
When acquiring live-cell imaging with fluorophores, it is crucial to select the appropriate wavelength range. Optimal excitation power, depletion laser power, pixel size, line accumulations, and dwell time should be empirically determined to maximize the signal-to-noise ratio (SNR) while minimizing photobleaching, particularly during extended time-lapse imaging. We also recommend positioning the cells in the center of the field of view (FOV), as the imaging quality is best at the center of the FOV.
Excitation power: Select an appropriate excitation wavelength based on the dye properties and experimental requirements. For HBmito Crimson, use an excitation wavelength of 640 nm. Start with a low excitation intensity (1%–5%) and gradually increase the excitation intensity in increments of 1%–3%. Determine an appropriate excitation power value based on the signal intensity observed in the image. Be cautious to avoid over-excitation, which can lead to photobleaching of the sample and prevent signal oversaturation.
Depletion laser power: Select an appropriate depletion wavelength. For HBmito Crimson, use a STED depletion laser wavelength of 775 nm. Start with a depletion laser power of 15%–20% (approximately 20–30 mW) and gradually increase the intensity in increments of 1%–3% (approximately 2–5 mW). Adjust the depletion light intensity based on the observed resolution and SNR in the images.
Pixel size: In STED microscopy, setting an appropriate pixel size is crucial to satisfy the Nyquist sampling theorem and ensure that the acquired images accurately reflect the sample's details and resolution. Pixel size should be at least 1/2 of the resolution, i.e., pixel size ≤ d/2.
Dwell time: Dwell time determines the exposure time for each pixel, thereby influencing the SNR and resolution of the image. For most samples, the setting of dwell time is typically between 1 and 10 μs. This range ensures that the image maintains an adequate SNR. If the sample signal is relatively weak, the exposure time can be increased to 10–20 μs to enhance the signal collection per pixel. A longer dwell time will improve the SNR, but it also increases the overall imaging time and the risk of photobleaching.
Line accumulations: The setting of line accumulations affects the SNR and resolution of the image. For common samples, the default line accumulations are typically set between 2 and 4 times. This range significantly enhances the SNR while avoiding excessive exposure that could lead to photobleaching. If the SNR is weak, the number of line accumulations can be increased, ensuring the sample remains stable and motionless during the acquisition process and does not shift due to prolonged acquisition times. For unstable samples, it is advisable to reduce the number of line accumulations. For experiments requiring high resolution and high SNR, consider setting line accumulations to higher values (e.g., 8 or more). Conversely, for rapid imaging needs or samples prone to photobleaching, lower line accumulations (e.g., 1–2 times) can be selected to reduce photobleaching and imaging time.
To determine the optimal imaging parameters, we recommend using the method of controlled variables, systematically altering one parameter at a time to observe its effect on image quality. This approach allows for achieving the best image quality and highest resolution.
1. Power on the microscope system, ensuring proper communication between the software and hardware, and open the Abberior Facility Line software.
2. Select the 63× oil immersion objective and apply immersion oil to the lens.
3. Place the standard beads sample on the sample holder, open the software, select the alignment function, and wait for the system dialog to display the message "alignment successful," indicating the program is complete. Ensure both excitation and depletion lasers are properly aligned.
4. Connect the live-cell incubation chamber to the microscope system. Once the environmental conditions in the incubation chamber stabilize at 37 °C, 5% CO2, and 75% humidity, add the prepared sample.
5. Use the microscope’s autofocus function to locate the sample’s focal plane. The Abberior autofocus actively stabilizes the Z-position of the sample, with continuous Z-drift compensation ensuring that focal drift does not occur during confocal and STED imaging.
6. Identify a suitable imaging area in confocal mode.
7. Adjust the imaging parameters accordingly.
As shown in Figure 2, λEx 640 nm: 15.7 μW; λDep 775 nm: 71.2 mW; pixel size: 20 nm; line accumulations: 3; dwell time: 6.5 μs; ROI: 44.5 μm × 50.74 μm; detection wavelength: 650–750 nm.
Figure 2. Comparison of confocal and STED imaging results in living COS7 cell mitochondria labeled with HBmito Crimson. Scale bar in the original image is 5 μm. Scale bar in the enlarged image is 1 μm.
8. Acquire STED images, ensuring that confocal and STED images are collected simultaneously.
9. For long-term imaging, appropriately reduce the excitation power, depletion power, line accumulations, and dwell time based on empirical observations. Depletion laser intensity can be reduced to approximately half of that used for single-frame imaging to meet the imaging requirements. Set the time intervals and number of imaging frames, then acquire images.
As shown in Figure 3, λEx 640 nm: 12.75 μW; λDep 775 nm: 34.7 mW; pixel size: 25 nm; line accumulations: 3; dwell time: 5 μs; frame time: 8.22 s; ROI: 4.75 μm × 7.025 μm; detection wavelength: 650–750 nm.
Figure 3. Long-term imaging of mitochondria in COS7 cells labeled with HBmito Crimson; white arrow indicates fusion event. Scale bar: 1 μm.
Note: When determining parameters for long-term imaging, ensure the minimal necessary laser intensity and exposure time to achieve only the required resolution. For mitochondrial cristae imaging, the minimum resolution requirement is defined as the clear visualization of cristae structures at the given power level. By utilizing these optimized settings, long-term imaging can minimize photobleaching effects, allowing for the acquisition of more frames and extending the imaging duration.
10. Save the data.
D. Extended image processing
Process single-frame images directly using Huygens software, setting post-processing parameters for each channel separately. For long-term imaging, it is recommended to first perform intensity correction, followed by batch deconvolution in Huygens software.
1. Open the Huygens Professional software. Import the image files that need deconvolution by selecting File > Open.
2. Access the image properties by selecting the image in the main window. This option can be found under the Edit menu or by right-clicking the image and selecting Properties. Set the microscope parameters accordingly:
Objective lens: Enter the numerical aperture (N.A.) and magnification of the objective lens used to acquire the images.
Excitation wavelength (λEx): Specify the excitation wavelength used during imaging.
Emission wavelength (λEm): Enter the emission wavelength corresponding to the fluorescence channel.
Voxel size: Input the pixel size (in x, y, and z dimensions) used during image acquisition.
Refractive index: Enter the refractive index of the immersion medium used (e.g., oil, water, glycerol).
Pinhole radius: If using a confocal microscope, specify the pinhole radius or size.
3. Select point spread function (PSF): In the Restoration menu, select the PSF option. If the software does not automatically detect the PSF, you can manually select or load the appropriate PSF file. Select Deconvolution Wizard from the Restoration menu. Deconvolution is typically performed using the classic maximum likelihood estimation (CMLE) algorithm, where the software automatically calculates parameters like SNR and background. These default settings are optimized to handle most images. However, parameters such as iteration count, SNR, and background can be manually adjusted to enhance image quality, especially for more complex or low-SNR images.
4. Click the Run button to start deconvolution processing.
5. After processing, use the software's visualization tool to view the deconvolved image.
6. Save the processed image by selecting Save As from the File menu. Save the image in common formats such as TIFF, JPEG, or other available formats.
7. Use the Batch Processor tool to perform batch deconvolution processing on multiple image files. After setting the batch processing parameters and file path, click the Start Batch button to start processing.
Data analysis
The resolution of the STED images is analyzed based on the full width at half maximum (FWHM) of mitochondrial structures. This analysis provides a quantitative measure of the system's resolution, allowing us to accurately assess the spatial resolution achieved during imaging.
1. Launch ImageJ software and open the desired image by selecting File > Open.
2. Duplicate the field of image to be analyzed. Use the Straight Line function in ImageJ to select multiple cristae within this area, as shown in Figure 4, and generate the signal intensity profile at the specified location following the direction indicated by the arrows.
Figure 4. Enlarged STED and STED+ results and fluorescence signal intensity profile correspond to the white arrow. Scale bar: 1 μm. Results were processed with the commercial deconvolution software Huygens (SVI, Netherlands).
3. Utilize the Analyze > Plot Profile function in ImageJ to obtain the signal intensity profile. Click List to open the intensity values, then copy these values into Origin software.
4. In Origin, apply a Gaussian Fit to derive the Gaussian fitting curve, as shown in Figure 5.
5. From the Gaussian fit, calculate the standard deviation (σ). Use the formula FWHM = 2.355 × σ to determine the FWHM of the image.
Figure 5. STED and STED+ results and the fitted fluorescence signal intensity distribution correspond to the white arrows. Scale bar: 1 μm.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article:
Ren et al [11]. Visualization of cristae and mtDNA interactions via STED nanoscopy using a low saturation power probe. (Figures 2–6).
General notes and troubleshooting
General notes
1. The dye concentration must be carefully optimized; neither too high nor too low. The recommended dye concentration of HBmito Crimson is 500 nM and the dyeing time is 10 min. Adjustments can be made according to the specific experimental requirements.
2. Select an appropriate laser wavelength for excitation based on the specific dye.
3. Calibrate the excitation and depletion lasers of the STED system to achieve optimal STED performance.
4. Maintaining stable laboratory temperature and humidity is essential to avoid environmental fluctuations that could impact microscope performance.
Troubleshooting
Problem 1: Motion artifacts in STED imaging
Possible cause: Sample jitter or mitochondrial movement.
Solution: It is essential to ensure the sample remains stable and motionless during imaging. In live-cell imaging, slow imaging speeds can lead to mitochondrial movement, resulting in motion artifacts. The imaging speed is limited by pixel dwell time and the line accumulations. Therefore, it is important to minimize line accumulations and dwell time while ensuring sufficient fluorescence signal collection. Additionally, reducing the FOV can help increase imaging speed and reduce the impact of mitochondrial movement.
Problem 2: Insufficient resolution in STED images
Possible cause: Improper alignment of the STED optical path or suboptimal parameter settings.
Solution: Ensure that the system is correctly calibrated. After calibration, image standard 40 nm beads. Perform Gaussian fitting on the signal intensity of individual beads to obtain their FWHM. If the FWHM reaches 40 nm, the system is in good condition. During sample imaging, improper parameter settings can lead to insufficient resolution. Refer to the parameter setting recommendations in the procedure. When adjusting parameters, avoid repeatedly imaging the same FOV to minimize photobleaching effects.
Acknowledgments
This protocol is related to the following paper: Ren et al. Visualization of cristae and mtDNA interactions via STED nanoscopy using a low saturation power probe. DOI: 10.1038/s41377-024-01463-9. This work was supported by the National Key R&D Program of China (2022YFC3401100), National Natural Science Foundation of China (22177024, 62025501, 31971376, 92150301), and Central Guidance Fund for Local Science and Technology Development (246Z1302G). We thank the National Center for Protein Sciences at Peking University in Beijing, China, for assistance with STED super-resolution imaging. We thank Abberior China and Optofem Technology Limited for providing Facility Line STED and Huygens software.
Competing interests
B.G. is an inventor of the awarded patent (ZL202011401937.1) of HBmito Crimson. The other authors declare no competing interests.
Ethical considerations
There are no ethical considerations associated with this protocol.
References
Spinelli, J. B. and Haigis, M. C. (2018). The multifaceted contributions of mitochondria to cellular metabolism. Nat Cell Biol. 20(7): 745–754. https://doi.org/10.1038/s41556-018-0124-1
Tan, J. X. and Finkel, T. (2020). Mitochondria as intracellular signaling platforms in health and disease. J Cell Biol. 219(5): e202002179. https://doi.org/10.1083/jcb.202002179
Rizzuto, R., De Stefani, D., Raffaello, A. and Mammucari, C. (2012). Mitochondria as sensors and regulators of calcium signaling. Nat Rev Mol Cell Biol. 13(9): 566–578. https://doi.org/10.1038/nrm3412
Lill, R., Hoffmann, B., Molik, S., Pierik, A. J., Rietzschel, N., Stehling, O., Uzarska, M. A., Webert, H., Wilbrecht, C., and Mühlenhoff, U. (2012). The role of mitochondria in cellular iron-sulfur protein biogenesis and iron metabolism. Biochim Biophys Acta. 1823(9): 1491–1508. https://doi.org/10.1016/j.bbamcr.2012.05.009
Friedman, J. R. and Nunnari, J. (2014). Mitochondrial form and function. Nature. 505(7483): 335–343. https://doi.org/10.1038/nature12985
Baker, N., Patel, J. and Khacho, M. (2019). Linking mitochondrial dynamics, cristae remodeling and supercomplex formation: How mitochondrial structure can regulate bioenergetics. Mitochondrion. 49: 259–268. https://doi.org/10.1016/j.mito.2019.06.003
Cogliati, S., Enriquez, J. A. and Scorrano, L. (2016). Mitochondrial Cristae: Where Beauty Meets Functionality. Trends Biochem Sci. 41(3): 261–273. https://doi.org/10.1016/j.tibs.2016.01.001
Kondadi, A. K., Anand, R., and Reichert, A. S. (2020). Cristae Membrane Dynamics-A Paradigm Change. Trends Cell Biol. 30(12): 923–936. https://doi.org/10.1016/j.tcb.2020.08.008
Liu, T., Stephan, T., Chen, P., Keller-Findeisen, J., Chen, J., Riedel, D., Yang, Z., Jakobs, S. and Chen, Z. (2022). Multi-color live-cell STED nanoscopy of mitochondria with a gentle inner membrane stain. Proc Natl Acad Sci USA. 119(52): e2215799119. https://doi.org/10.1073/pnas.2215799119
Stephan, T., Roesch, A., Riedel, D. and Jakobs, S. (2019). Live-cell STED nanoscopy of mitochondrial cristae. Sci Rep. 9(1): e1038/s41598–019–48838–2. https://doi.org/10.1038/s41598-019-48838-2
Ren, W., Ge, X., Li, M., Sun, J., Li, S., Gao, S., Shan, C., Gao, B. and Xi, P. (2024). Visualization of cristae and mtDNA interactions via STED nanoscopy using a low saturation power probe. Light Sci Appl. 13(1): 116. https://doi.org/10.1038/s41377-024-01463-9
Article Information
Publication history
Received: Aug 11, 2024
Accepted: Oct 27, 2024
Available online: Nov 27, 2024
Published: Jan 5, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
How to cite
Category
Biophysics > Biophotonics
Cell Biology > Cell imaging > Super resolution imaging
Cell Biology > Cell staining > Organelle
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TS Tadashi Suzuki
Published: Vol 15, Iss 1, Jan 5, 2025
DOI: 10.21769/BioProtoc.5151 Views: 202
Reviewed by: Laxmi Narayan MishraFNU Priyanka Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in The Journal of Biological Chemistry Apr 2024
Abstract
Cytosolic peptide: N-glycanase (PNGase/NGLY1 in mammals), an amidase classified under EC:3.5.1.52, is a highly conserved enzyme across eukaryotes that catalyzes the removal of N-glycans from glycoproteins, converting N-glycosylated asparagine residues into aspartic acid. This enzyme also plays a role in the quality control system for nascent glycoproteins. Despite the development of non-radioisotope-based assay systems such as those using S-alkylated RNase or fluorescent-labeled glycopeptides as substrates, these methods are incompatible with crude enzyme sources, primarily due to the degradation of reaction products by contaminating endogenous proteases. We previously developed an assay system using a 5-carboxyfluorescein-labeled glycosylated cyclo-heptapeptide (5FAM-GCP), a substrate remarkably resistant to endogenous peptidase activity. This system enables the accurate measurement of endogenous NGLY1 activity in various samples, including cell lines, tissues, peripheral blood mononuclear cells, and NGLY1-deficient patient-derived cells, without the interference of proteolytic degradation. We recently advanced this approach by producing a novel fluorescence resonance energy transfer (FRET)-based GCP probe (fGCP) and demonstrated its ability to detect endogenous NGLY1 activity across diverse enzyme sources via fluorescence on multiarray plates. This innovative and straightforward assay now offers reliable disease diagnostics and also allows the measurement of endogenous PNGase/NGLY1 activities across various organisms.
Key features
• fGCP assay enables measurement of endogenous PNGase/NGLY1 activity in cells and tissues.
• An aliquot of 1–5 × 106 cells or 50–100 μg of protein extract from tissues is used for this assay.
• This assay enables microplate-based real-time measurement of endogenous PNGase/NGLY1 activities.
• This protocol requires a fluorescence plate reader equipped with an incubation function.
Keywords: Peptide:N-glycanase NGLY1 FRET Enzyme assay Glycosylated cyclopeptide Microplate-based assay
Graphical overview
Overview of the real-time measurement of endogenous PNGase/NGLY1 activities in tissues/cells using fGCP assay
Background
Cytosolic peptide: N-glycanase (PNGase; NGLY1 in mammals) is an amidase that catalyzes the removal of N-glycans from the consensus sequences (Asn-Xaa-Ser/Thr, where Xaa represents any amino acid except proline) of glycoproteins, converting N-glycosylated Asn into Asp residues through deglycosylation [1]. Cytosolic PNGase also plays a role in the quality control system for newly synthesized glycoproteins [2].
To explore the biochemical properties of this enzyme, 14C-labeled glycopeptides, such as pentapeptides that carry asialoglycans derived from fetuin, have been commonly used as substrates for measuring PNGase activity [3]. Enzyme activity was measured based on the radioactivity of the reaction products, which were separated by paper chromatography or paper electrophoresis. However, the preparation of radioisotope-labeled glycopeptides and the use of radioactive molecules make it difficult to perform this assay in standard laboratory settings, especially under the tight regulations for the use of radioactive isotopes in Japan. Although non-radioisotope-based assays have been developed (e.g., assays using S-alkylated RNase [4] or fluorescent-labeled glycopeptides [5]), they are incompatible with crude enzyme sources due to the degradation of reaction products by contaminating endogenous proteases. Therefore, there exists a need to develop an alternative, easy-to-handle assay method.
An autosomal recessive disorder linked to NGLY1, known as NGLY1 deficiency or congenital disorder of deglycosylation (NGLY-CDDG) [OMIM: 615273], was first reported in 2012 [6]. Since then, more than 100 patients have been identified worldwide, including in Europe, America, Australia, India, China, and Japan [7,8]. The disease exhibits a broad spectrum of symptoms, including global developmental delay and/or intellectual disability, abnormal EEG, seizures, movement disorders, hypolacrima or alacrima, and liver dysfunction [8–13]. Unfortunately, there are no effective treatments; however, recent studies have demonstrated that administering an adeno-associated viral vector serotype 9 carrying the human NGLY1 gene to Ngly1-deficient model rats aged 3 or 5 to 7 weeks through intracerebroventricular injection significantly improved their motor function defects [14–16]. Considering the importance of the therapeutic time window for gene therapies, early intervention may be crucial to alleviate the various symptoms caused by the dysfunction of the central nervous system in this disease. Therefore, there exists an urgent need for methods to enable the early diagnosis of NGLY1 deficiency by measuring endogenous NGLY1 activity in specimens from potential disease candidates.
A method for measuring endogenous NGLY1 activity using 5-carboxyfluorescein-labeled glycosylated cyclo-heptapeptide (5FAM-GCP) has been established previously [3,17,18]. This approach enables detecting endogenous PNGase/NGLY1 activities from various enzyme sources without the proteolytic degradation of reaction products during incubation with crude enzyme preparations. However, it requires HPLC for the separation and detection of products, which is often unavailable in standard clinical laboratories. Hence, it is crucial to develop a facile, sensitive probe for enzyme assay similar to MM3D, a fluorescence and quencher-based FRET probe designed for detecting ENGase activity [19]. We recently developed a novel FRET-based GCP probe (fGCP) consisting of a glycan modified with a fluorophore-labeled bisected-GlcNAc [aminomethylcoumarin acetate-labeled GlcNAc (AMCA-GlcNAc)] and a cyclo-heptapeptide modified with a quencher, 4-((4-(dimethylamino)phenyl)azo)benzoic acid (Dabcyl) (Figure 1) [20]. This method allows the detection of endogenous NGLY1 activity in various enzyme sources via fluorescence on multiarray plates. Our novel assay method could provide a reliable diagnostic tool and valuable insights into the regulation of PNGase/NGLY1 activities in various organisms.
Figure 1. Structure of 5-carboxyfluorescein- and dabcyl-labeled glycosylated cyclo-heptapeptide (fGCP) and deglycosylation reaction catalyzed by PNGase [20]. The bisected-GlcNAc and lysine residues on the glycosylated cyclo-heptapeptide were labeled with a fluorophore, aminomethylcoumarin acetate (AMCA), and a quencher, 4-((4-(dimethylamino)phenyl)azo)benzoic acid (Dabcyl), respectively. The right bracket illustrates a schematic of fGCP and deglycosylation reaction catalyzed by PNGase/NGLY1.
Materials and reagents
Biological materials
1. Rat brain tissues (a rat outbred strain, Sprague-Dawley)
2. Cell lines (e.g., HeLa and HEK293 cells)
3. Fibroblast derived from healthy subjects and NGLY1-deficiency patients (available from Coriell.org)
Reagents
1. 5-Carboxyfluorescein- and dabcyl-labeled glycosylated cyclo-heptapeptide (fGCP, Mw: 2924.9) (GlyTech, Inc.)
2. Sucrose (FUJIFILM Wako Chemicals, catalog number: 196-00015)
3. EDTA (FUJIFILM Wako Chemicals, catalog number: 345-01865)
4. Trizma base (Sigma-Aldrich, catalog number: T1503)
5. NP-40 (IGEPAL® CA-630) (MPBIO, catalog number: 198596)
6. Hydrochloric acid (HCl) (FUJIFILM Wako Chemicals, catalog number: 080-01066)
7. cOmplete EDTA-free protease inhibitor cocktail (Merck-Millipore, catalog number: 11836170001)
8. Pefabloc SC (Merck-Millipore, catalog number: 11429868001)
9. Rabeprazole sodium salt (Tokyo Chemical Industry Co. Ltd., catalog number: R0115)
10. Dithiothreitol (DTT) (FUJIFILM Wako Chemicals, catalog number: M02712)
11. Powermasher II (Nippi-Inc., catalog number: 891-300)
12. Biomasher II (1.5 mL tube) (Nippi-Inc., catalog number: 320-103)
Solutions
1. 10× NGLY1 buffer (see Recipes)
2. Lysis buffer for animal tissues (see Recipes)
3. Lysis buffer for cultured cells (see Recipes)
4. 1 mM fGCP stock solution (see Recipes)
5. 100 μM fGCP working solution (see Recipes)
Recipes
1. 10× NGLY1 buffer
Reagent Final concentration Quantity or Volume
1 M Tris-HCl (pH 7.5) 50 mM 5 mL
Sucrose 10 mM 342 mg
500 mM EDTA (pH 8.0) 5 mM 1 mL
Total n/a 100 mL
Store at room temperature. This buffer remains stable at room temperature for at least one year.
2. Lysis buffer for animal tissues
Reagent Final concentration Quantity or Volume
10× NGLY1 buffer 1× 100 μL
100 mM DTT 1 mM 10 μL
100 mM Pefabloc SC 1 mM 10 μL
50× protease inhibitor cocktail 1× 20 μL
5 mM Rabeprazole 50 μM 10 μL
Distilled water n/a 850 μL
Total n/a 1 mL
40 μL of lysis buffer is used for one reaction. The reagent should be prepared immediately before use.
3. Lysis buffer for cultured cells
Reagent Final concentration Quantity or Volume
10× NGLY1 buffer 1× 100 μL
10% (v/v) NP-40 0.5% (v/v) 50 μL
100 mM DTT 1 mM 10 μL
100 mM Pefabloc SC 1 mM 10 μL
50× protease inhibitor cocktail 1× 20 μL
5 mM Rabeprazole 50 μM 10 μL
Distilled water n/a 800 μL
Total n/a 1 mL
40 μL of the buffer is used for each reaction. The reagent should be prepared immediately before use.
4. 1 mM fGCP stock solution
Reagent Final concentration Quantity or Volume
fGCP 1 mM 1 mg
Distilled water n/a 342 μL
Store at -20 °C. This stock remains stable at -20 °C for at least 1–2 years.
5. 100 μM fGCP working solution
Reagent Final concentration Quantity or Volume
1 mM fGCP 100 μM 1 μL
Lysis buffer n/a 9 μL
Total n/a 10 μL
10 μL of the solution is used for each reaction.
Laboratory supplies
1. 96-well black polystyrene microplate (clear flat bottom) (Corning, catalog number: CLS3603)
Equipment
1. Refrigerated microcentrifuge
2. Sonicator (TOMY, model: UR-21P)
3. Varioskan LUX multimode microplate reader (Thermo, model: VL0000D0)
Procedure
A. Preparation of cell lysate from cultured cells
1. Collect cultured cells (5 × 106 cells) into a tube and wash them with PBS (see Note 1).
2. Resuspend the cells in 50 μL of lysis buffer for cultured cells.
3. Incubate the suspension on ice for 10 min to disrupt the cells.
4. Clarify the samples by centrifugation at 20,000× g for 5 min at 4 °C.
5. Transfer the supernatant to a new tube and use as the enzyme source (see Notes 2 and 3).
B. Preparation of cell lysate from rodent tissues
1. Transfer tissues of interest (e.g., 25–50 mg of the brain) to 1.5 mL Biomasher II tube (see Note 4).
2. Resuspend the tissue into 250 μL of NGLY1 buffer.
3. Lyse the tissue four times by homogenizing for 20 s followed by a cooling period of 20 s on ice using Powermasher II.
4. Clarify the samples by centrifugation at 20,000× g for 5 min at 4 °C.
5. Transfer the supernatant to a new tube and use as the enzyme source (see Note 3).
6. Calculate protein concentration in the cell lysate by protein assay (e.g., BCA assay) (see Note 5).
C. Real-time measurement of PNGase/NGLY1 activity
1. Transfer 40 μL of the enzyme source to a 96-well black polystyrene microplate (see Note 6).
2. Add 10 μL of 100 μM fGCP solution to the well loaded with the enzyme source and mix the solution by pipetting.
3. Immediately set the plate on the Varioskan LUX multimode microplate reader (see Note 7).
4. Measure fluorescence intensity every 15 min with incubation at 25 °C for 6–12 h (λ excitation: 353 nm; λ emission: 450 nm) (Figure 2A and C, Figure 3A and C) (see Notes 8 and 9).
Figure 2. Real-time measurement of NGLY1 activity in cultured cell lines and evaluation of the endogenous NGLY1 activity in patients’ fibroblast using fGCP assay [20]. A. Analysis of endogenous NGLY1 activity in HeLa and HEK293 cells. NGLY1-KO cells show no endogenous NGLY1 activity in both cell lines. Each cell extract prepared from 5 × 106 cells was incubated with fGCP for 12 h at 25 °C. The fluorescence intensity of fGCP was measured every 15 min. B. Two parameters, initial slope (0–90 min) and maximum fluorescence intensity of endogenous NGLY1 activity were measured in HeLa and HEK293 cells. C. Real-time measurement of endogenous NGLY1 activity (measured every 15 min) in fibroblasts. The fluorescence intensity of each sample was measured by incubating fGCP with crude cell extract prepared from 5 × 106 cells for 8 h at 25 °C. D. Quantitative analysis of endogenous NGLY1 activity in fibroblasts. The horizontal line represents the means of biological triplicates for control- or patient-derived samples. Error bars are mean ± S.D. from biological triplicates. For statistical analysis, a Student’s t-test was applied. *, **, and *** represent p < 0.05, p < 0.01, and p < 0.001, respectively.
Figure 3. Real-time measurement of NGLY1 activity in rat tissues: liver and brain [20]. A. Real-time measurement of endogenous Ngly1 activity (measured every 15 min) in rat liver of various ages. The fluorescence intensity of each sample was measured by incubating fGCP with 116 μg of protein extract from the liver for 12 h at 25 °C. B. Quantitative analysis of (A). C. Real-time measurement of endogenous Ngly1 activity (measured every 15 min) in rat brain. Fluorescence intensity of each sample was measured by incubating fGCP with 116 μg of protein extract from the brain for 12 h at 25 °C. D. Quantitative analysis of C. Error bars are means ± S.D. (n = 3 rats in each age). For statistical analysis, a Student’s t-test was applied. *, **, and *** represent p < 0.05, p < 0.01, and p < 0.001, respectively.
Data analysis
1. For calculating the fluorescence intensity in each sample, the blank value should be subtracted from all other sample values.
2. Fluorescence intensity should be normalized by cell number or protein concentration.
3. Reaction curves can be evaluated using the initial slope (slope of 0–90 min) and the maximum intensity of the reaction (Figure 2B and D, Figure 3B and D) (see Notes 10 and 11).
Validation of protocol
This protocol has been used and validated in the following research article:
• Hirayama et al. [20]. Development of a fluorescence and quencher-based FRET assay for detection of endogenous peptide: N-glycanase/NGLY1 activity. J Biol Chem 300(4): 107121 (Figures 4–6).]
General notes and troubleshooting
General notes
1. The collected cells can be flash frozen in liquid nitrogen and stored at -80 °C until use.
2. For one assay, 1–5 × 106 cells were required to prepare the crude extract.
3. Cells or tissue lysates should be prepared immediately before the assay, as NGLY1 activity is drastically decreased in frozen crude lysates.
4. The preserved frozen tissues (stored at -80 °C) are also used as a source of the enzyme in this assay.
5. Lysates containing 50–100 μg of proteins are required for one assay.
6. Prepare a blank well containing 40 μL of the same buffer as samples (lysis buffer for animal tissues or cultured cells) and 10 μL of 100 μM fGCP working solution.
7. Keep the lid of the assay plate closed to prevent evaporation of the solution.
8. For the evaluation of endogenous NGLY1 activity, it is preferable to perform assays of the lysates prepared from NGLY1-KO cells or lysates treated with 100 μM of zVAD-fmk, a potent NGLY1-inhibitor [21], as a negative control.
9. Replicate experiments (biological or technical) should be carried out to ensure accurate interpretation of the results through statistical analysis.
10. It is possible to determine the initial slope by measuring two time points (0 and 90 min) to compare the relative enzyme activity among the samples. However, real-time measuring of the reaction for 8–12 h is preferable to obtain more information on the enzyme properties (e.g., background level, maximum activity, and time to reach plateau)
11. To calculate the specific activity of NGLY1 (expressed as pmol of deglycosylated fGCP per minute normalized by protein concentration; pmol/min/mg protein), create a standard curve of the concentration of fGCP treated with PNGase F and the fluorescence intensity. Then, calculate the pmol of the product (i.e., deglycosylated substrate) from the standard curve. Alternatively, the 5FAM-GCP assay [18], which can measure enzyme activity through the separation of the substrate and product using HPLC, can also be used.
Troubleshooting
Problem 1: Fluorescence signal is low throughout the experiment.
Possible cause: Number of cells/total tissues for the preparation of cell lysates is too low.
Solution(s): Prepare more cells/tissues.
Acknowledgments
Research reported in this publication was supported by the T-CiRA Program (NGLY1 Deficiency Project), the RIKEN Pioneering Project (“Glyco-lipidologue Initiative”), Japan Agency for Medical Research and Development Core Research for Evolutional Science and Technology (AMED-CREST) Grant JP24gm14100003h0005 (to T.S.), and by KAKENHI Grant Number JP22K06155 (to H.H.) from the Ministry of Education, Culture, Sports, Science, and Technology (MEXT)/Japan Society for the Promotion of Science (JSPS). The image of cells in the Graphical overview was provided by DBCLS Togo Picture Gallery (2016 DBCLS TogoTV; https://togotv.dbcls.jp/pics.html). We thank Enago (https://www.enago.jp) for the English language review. This protocol was adapted and modified from Hirayama et al. [20].
Competing interests
The authors declare that they have no competing interests.
Ethical considerations
Care procedures and experiments of a rat outbred strain, Sprague–Dawley, conformed to the association for assessment and accreditation of laboratory animal care guidelines. All experiments using animals were approved by the experimental animal care and use committee of our organization.
References
Suzuki, T., Park, H. and Lennarz, W. J. (2002). Cytoplasmic peptide:N-glycanase (PNGase) in eukaryotic cells: occurrence, primary structure, and potential functions. FASEB J. 16(7): 635–641.
Hirayama, H., Hosomi, A. and Suzuki, T. (2015). Physiological and molecular functions of the cytosolic peptide:N-glycanase. Semin Cell Dev Biol. 41: 110–120.
Hirayama, H. and Suzuki, T. (2022). Assay for the peptide:N-glycanase/NGLY1 and disease-specific biomarkers for diagnosing NGLY1 deficiency. J Biochem. 171(2): 169–176.
Suzuki, T. (2005). A simple, sensitive in vitro assay for cytoplasmic deglycosylation by peptide: N-glycanase. Methods. 35(4): 360–365.
Taga, E. M., Waheed, A. and Van Etten, R. L. (1984). Structural and chemical characterization of a homogeneous peptide N-glycosidase from almond. Biochemistry. 23(5): 815–822.
Need, A. C., Shashi, V., Hitomi, Y., Schoch, K., Shianna, K. V., McDonald, M. T., Meisler, M. H. and Goldstein, D. B. (2012). Clinical application of exome sequencing in undiagnosed genetic conditions. J Med Genet. 49(6): 353–361.
Pandey, A., Adams, J. M., Han, S. Y. and Jafar-Nejad, H. (2022). NGLY1 Deficiency, a Congenital Disorder of Deglycosylation: From Disease Gene Function to Pathophysiology. Cells. 11(7): 1155.
Abuduxikuer, K., Zou, L., Wang, L., Chen, L. and Wang, J. S. (2020). Novel NGLY1 gene variants in Chinese children with global developmental delay, microcephaly, hypotonia, hypertransaminasemia, alacrimia, and feeding difficulty. J Hum Genet. 65(4): 387–396.
Enns, G. M., Shashi, V., Bainbridge, M., Gambello, M. J., Zahir, F. R., Bast, T., Crimian, R., Schoch, K., Platt, J., Cox, R., et al. (2014). Mutations in NGLY1 cause an inherited disorder of the endoplasmic reticulum-associated degradation pathway. Genet Med. 16(10): 751–758.
Ge, H., Wu, Q., Lu, H., Huang, Y., Zhou, T., Tan, D. and ZhongqinJin (2020). Two novel compound heterozygous mutations in NGLY1 as a cause of congenital disorder of deglycosylation: a case presentation. BMC Med Genet. 21(1): 135.
Lam, C., Ferreira, C., Krasnewich, D., Toro, C., Latham, L., Zein, W. M., Lehky, T., Brewer, C., Baker, E. H., Thurm, A., et al. (2017). Prospective phenotyping of NGLY1-CDDG, the first congenital disorder of deglycosylation. Genet Med. 19(2): 160–168.
Lipari Pinto, P., Machado, C., Janeiro, P., Dupont, J., Quintas, S., Sousa, A. B. and Gaspar, A. (2020). NGLY1 deficiency-A rare congenital disorder of deglycosylation. JIMD Rep. 53(1): 2–9.
Sonoda, Y., Fujita, A., Torio, M., Mukaino, T., Sakata, A., Matsukura, M., Yonemoto, K., Hatae, K., Ichimiya, Y., Chong, P. F., et al. (2024). Progressive myoclonic epilepsy as an expanding phenotype of NGLY1-associated congenital deglycosylation disorder: A case report and review of the literature. Eur J Med Genet. 67: 104895.
Asahina, M., Fujinawa, R., Hirayama, H., Tozawa, R., Kajii, Y. and Suzuki, T. (2021). Reversibility of motor dysfunction in the rat model of NGLY1 deficiency. Molecular Brain. 14(1): 91.
Fujihira, H., Asahina, M. and Suzuki, T. (2022). Physiological importance of NGLY1, as revealed by rodent model analyses. J Biochem. 171(2): 161–167.
Zhu, L., Tan, B., Dwight, S. S., Beahm, B., Wilsey, M., Crawford, B. E., Schweighardt, B., Cook, J. W., Wechsler, T. and Mueller, W. F. (2022). AAV9-NGLY1 gene replacement therapy improves phenotypic and biomarker endpoints in a rat model of NGLY1 Deficiency. Mol Ther Methods Clin Dev. 27: 259–271.
Hirayama, H., Tachida, Y., Seino, J. and Suzuki, T. (2022). A method for assaying peptide: N-glycanase/N-glycanase 1 activities in crude extracts using an N-glycosylated cyclopeptide. Glycobiology. 32(2): 110–122.
Hirayama, H. and Suzuki, T. (2022). Enzyme assay of endogenous activity of peptide-N-glycanase (PNGase)/N-Glycanase 1 (NGLY1) in cells and tissues using glycosylated cyclopeptide as a substrate. In: Nishihara, S., Angata, K., Aoki-Kinoshita, K. F. and Hirabayashi, J. (Eds.) Glycoscience Protocols (GlycoPODv2).
Ishii, N., Muto, H., Nagata, M., Sano, K., Sato, I., Iino, K., Matsuzaki, Y., Katoh, T., Yamamoto, K. and Matsuo, I. (2023). A fluorogenic probe for core-fucosylated glycan-preferred ENGase. Carbohydr Res. 523: 108724.
Hirayama, H., Tachida, Y., Fujinawa, R., Matsuda, Y., Murase, T., Nishiuchi, Y. and Suzuki, T. (2024). Development of a fluorescence and quencher-based FRET assay for detection of endogenous peptide:N-glycanase/NGLY1 activity. J Biol Chem. 300(4): 107121.
Misaghi, S., Pacold, M. E., Blom, D., Ploegh, H. L. and Korbel, G. A. (2004). Using a small molecule inhibitor of peptide: N-glycanase to probe its role in glycoprotein turnover. Chem Biol. 11(12): 1677–1687.
Article Information
Publication history
Received: Aug 25, 2024
Accepted: Nov 6, 2024
Available online: Nov 20, 2024
Published: Jan 5, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Biochemistry > Protein > Quantification
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Isolation of Viral Biofilms From HTLV-1 Chronically Infected T Cells and Integrity Analysis
CA Coline Arone
HD Hélène Dutartre
DM Delphine Muriaux
Published: Vol 14, Iss 24, Dec 20, 2024
DOI: 10.21769/BioProtoc.5152 Views: 269
Reviewed by: Alka MehraJibin SadasivanSrajan Kapoor
Download PDF
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Cited by
Original Research Article:
The authors used this protocol in mBio Oct 2023
Abstract
The human T-lymphotropic virus type-1 (HTLV-1) is an oncogenic retrovirus that predominantly spreads through cell-to-cell contact due to the limited infectivity of cell-free viruses. Among various modes of intercellular transmission, HTLV-1 biofilms emerge as adhesive structures, polarized at the cell surface, which encapsulate virions within a protective matrix. This biofilm is supposed to facilitate simultaneous virion delivery during infection. Yet, the molecular and functional intricacies of viral biofilms remain largely unexplored, despite their pivotal role in understanding retroviral pathogenesis. In this study, we optimized a protocol to isolate HTLV-1 biofilms from chronically infected T cells, facilitating their structural and molecular characterization using proteomic and super-resolution microscopy analyses. This protocol involves cultivating HTLV-1 chronically infected T cells at high density to facilitate the natural detachment of viral biofilms into the supernatant. Then, employing successive centrifugations, the cells are separated from the detached biofilms, and these structures are pelleted at medium speed (10,000× g). This method circumvents the need for mechanical, chemical, or enzymatic biofilm detachment, bypasses the use of ultracentrifugation, and enables us to resuspend the biofilms in the appropriate buffer for subsequent analyses such as western blotting or super-resolution microscopy imaging as presented.
Key features
• Isolation of viral biofilms from HTLV-1 chronically infected T cells after 4 days of culture at high cellular density.
• Structural analysis of viral biofilms using super-resolution microscopy techniques.
• Experiments performed in vitro within a confined biosafety level 3 (BSL3) environment.
• This protocol requires at least five days to complete.
Keywords: HTLV-1 Chronically infected T cells Viral biofilms Gag Env Super-resolution STED microscopy
Graphical overview
Fluorescent HTLV-1 viruses embedded in a biofilm
Background
The human T-lymphotropic virus type-I (HTLV-1) is an oncogenic retrovirus estimated to infect 5–10 million people worldwide [1]. Infected cells are characterized by the integration of the reverse-transcribed viral genome (proviral DNA) in the host genome, which can be expressed using the host transcription machinery, generating new virions and thus leading to chronic infection. Although most people remain asymptomatic following HTLV-1 exposure, chronic infections can lead to aggressive pathologies such as adult T-cell leukemia [2] or progressive inflammatory disorders like HTLV-1-associated myelopathy [3]. In vivo, HTLV-1 is primarily detected in CD4(+) T cells, and its dissemination among individuals occurs through three main routes: mother to child during breastfeeding, sexual contact, or exposure to HTLV-1-infected blood products [4]. Overall, this virus represents a major health issue as no existing strategy allows the efficient protection of exposed individuals. Therefore, fundamental research on HTLV-1 dissemination routes is essential to promote the expansion of pertinent therapeutic approaches.
Unlike most retroviruses, evidence strongly suggests that HTLV-1 cell-free virions are poorly infectious in vivo [5,6]. As such, HTLV-1 relies mostly on intercellular contacts to disseminate within its host. Three cell-to-cell transmission modes have been described: virological synapses, cellular conduits, and viral biofilms, which are of interest here [7]. Viral biofilms are cell-surface aggregates of viruses embedded in extracellular matrix components that accumulate virions at one or several poles of the infected T cells [8]. These structures allow the bulk delivery of infectious particles to target cells and may help to protect HTLV-1 virions from immune attacks [9,10]. While the relative importance of the different cell-to-cell transmission routes is difficult to ascertain in vivo, the removal of biofilms by heparin washes reduces the infectious capacity of HTLV-1-producing cells by 80% in vitro [9]. In addition, biofilms detached from HTLV-1-infected cells are infectious in vitro, as opposed to cell-free virions released in the supernatant of the same cell population [9,11], indicating that the transfer of HTLV-1 biofilms through cell-to-cell contact is the most efficient pathway to infect new cells.
To identify molecular factors involved in HTLV-1 biofilm architecture and explore their impact on viral transmission, we sought to isolate these structures, analyze their molecular composition using mass spectrometry, and confirm by super-resolution STED (stimulated-emission-depletion) microscopy [12]. Here, we will detail the protocol of biofilm isolation that we used in Arone et al. [13] and the techniques employed to validate our protocol (i.e., western blotting and STED microscopy). This protocol is an optimized version of procedures published in Pais-Correia et al. [9] and Alais et al. [11]. In these studies, authors used chemical or mechanical methods, respectively, to detach viral biofilms from HTLV-1 chronically infected T cells and used ultracentrifugation to collect them. While our protocol is also based on cell culture at high density to enrich viral biofilms in the supernatant, it circumvents the need for mechanical or chemical biofilm detachment and bypasses the use of ultracentrifugation. Finally, using immuno-staining coupled with super-resolution STED microscopy techniques [13], we confirm that our isolation protocol preserves the integrity of viral biofilms that contain unaltered virions.
Materials and reagents
Biological materials
HTLV-1 chronically infected T cells (C91-PL) derived from umbilical cord blood cells (CVCL_0197, https://www.cellosaurus.org/CVCL_0197).
pHTLV Gag-YFP plasmid was described previously in Heidecker et al. [14] and is a kind gift from former Dr. Derse’s lab.
Reagents
Phosphate buffered saline (PBS) pH 7,4 (Gibco, catalog number: 11503387)
Roswell Park Memorial Institute (RPMI) medium (Gibco, catalog number: 11875093)
Opti-MEM reduced serum medium (Gibco, catalog number: 31985062)
Fetal calf serum (FCS) (Sigma-Aldrich, catalog number: F7524)
Penicillin-streptomycin (Sigma-Aldrich, catalog number: P4333)
Trypan blue stain 0.4% (Logos biosystems, catalog number: T13001)
Poly-L-lysine hydrobromide, 5 mg (Sigma-Aldrich, catalog number: P6282)
Bovine serum albumin (BSA) lyophilized powder (Sigma-Aldrich, catalog number: A9418)
32% paraformaldehyde (PFA) without methanol (Thermo Fisher Scientific, catalog number: 047377.9L)
Triton X-100 (Sigma-Aldrich, catalog number: X100)
NaCl 5M solution (Thermo Fisher Scientific, catalog number: AM9760G)
NaCl powder (Sigma-Aldrich, catalog number: S9888)
EDTA 0.5M pH 8.0 (Thermo Fisher Scientific, catalog number: 15575020)
40% acrylamide/bisacrylamide (Euromedex, catalog number: EU0063-C)
Saponin (Sigma-Aldrich, catalog number: SAE0073)
NH4Cl (Sigma-Aldrich, catalog number: A9434)
Methanol (Honeywell, catalog number: 14262-1L)
37% hydrochloric acid (HCl) solution (Honeywell, catalog number: 30721)
Trizma base, crystalline (Sigma-Aldrich, catalog number: T1503)
Tris-glycine 10× buffer (Euromedex, catalog number: EU0550)
Tris-glycine SDS 10× buffer (Euromedex, catalog number: EU0510)
Tris/HCl 1M pH 7.4 (Thermo Fisher Scientific, catalog number: J67501.AE)
10% ammonium persulfate (APS) (Thermo Fisher Scientific, catalog number: 17874)
TEMED (Sigma-Aldrich, catalog number: T9281)
Laemmli SDS 4× buffer (Thermo Fisher Scientific, catalog number: J60015.AC)
Sterile Braun water (Dutscher, catalog number: 921120)
PageRuler prestained protein ladder (Thermo Fisher Scientific, catalog number: 26616)
Tween 20 (Thermo Fisher Scientific, catalog number: 85115)
Milk powder (Régilait)
ECL prime western blotting detection reagent (Amersham, catalog number: 10308449)
Anti-HTLV-1 Gagp19 primary mouse antibody (Zeptometrix, catalog number: 801108)
Anti-HTLV-1 Envgp46 primary mouse antibody (Zeptometrix, catalog number: 0801127)
Anti-YFP primary rabbit antibody (Thermo Fisher Scientific, catalog number: A-11122)
Anti-mouse IgG STAR Red (Abberior, catalog number: 52283)
Anti-rabbit IgG STAR 580 (Abberior, catalog number: 41367)
Anti-mouse IgG HRP (Dako, catalog number: P0260)
Solutions
TNE buffer (see Recipes)
Protein solubilization buffer (see Recipes)
Poly-L-lysine coating solution (see Recipes)
Fixation buffer (see Recipes)
Permeabilization and saturation buffer (see Recipes)
Tris-HCl 0.5M pH 6.8 (see Recipes)
Tris-HCl 1M pH 8.0 (see Recipes)
Tris-HCl 1.5M pH 8.8 (see Recipes)
Tris-buffered saline (TBS) (see Recipes)
Tris-buffered saline with Tween (TBST) (see Recipes)
10% separating electrophoresis gel (see Recipes)
4% stacking electrophoresis gel (see Recipes)
Western blot running buffer (see Recipes)
Western blot transfer buffer (see Recipes)
Recipes
TNE buffer
Reagent Final concentration Quantity or Volume
Tris/HCl 0.5M pH 7.4 10mM 200 μL
NaCl 5M 100mM 400 μL
EDTA 0.5M pH 8.0 1mM 40 μL
Milli-Q H2O Up to 20 mL
Total 20 mL
Note: Store at room temperature.
Protein solubilization buffer
Reagent Final concentration Quantity or Volume
Triton X-100 0.2% 2 mL
PBS pH 7.4 99.8% 8 mL
Total 100% 10 mL
Note: Store at 4 °C.
Poly-L-lysine coating solution
Reagent Final concentration Quantity or Volume
Poly-L-lysine hydrobromide 0.01% 5 mg
PBS pH 7.4 99.9% Up to 50 mL
Total 100% 50 mL
Note: Store at -20 °C as 2 mL aliquots to avoid freeze/thaw cycles.
Fixation buffer
Reagent Final concentration Quantity or Volume
32% paraformaldehyde 4% 10 mL
PBS pH 7.4 96% 70 mL
Total 100% 80 mL
Note: Store at -20 °C as 2 mL aliquots to avoid freeze/thaw cycles.
Permeabilization and saturation buffer
Reagent Final concentration Quantity or Volume
Saponin 0.05% 5 mg
BSA 3% 1.5 g
PBS pH 7.4 96,95% Up to 50 mL
Total 100% 50 mL
Note: Mix thoroughly to homogenize completely. Use immediately.
Tris-HCl 0.5M pH 6.8
Reagent Final concentration Quantity or Volume
Trizma base 30 g
HCl (Adjust to pH 6.8) ~ 9.6 mL
Milli-Q H2O Up to 500 mL
Total 500 mL
Note: HCl should be added in increments of 1–5 mL in 400 mL of H2O until the correct pH is reached. Allow the solution to mix completely before adding any more HCl. Complete to 500 mL with H2O. Store at room temperature.
Tris-HCl 1 M pH 8
Reagent Final concentration Quantity or Volume
Trizma base 242 g
HCl (Adjust to pH 8) ~ 50 mL
Milli-Q H2O Up to 2 L
Total 2 L
Note: HCl should be added in increments of 1–5 mL in 1.6 L of H2O until the correct pH is reached. Allow the solution to mix completely before adding any more HCl. Complete to 500 mL with H2O. Store at room temperature.
Tris-HCl 1.5 M pH 8.8
Reagent Final concentration Quantity or Volume
Trizma base 90.75 g
HCl (Adjust to pH 8.8) ~ 9 mL
Milli-Q H2O Up to 500 mL
Total 500 mL
Note: HCl should be added in increments of 1–5 mL in 400 mL of H2O until the correct pH is reached. Allow the solution to mix completely before adding any more HCl. Complete to 500 mL with H2O. Store at room temperature.
Tris-buffered saline (TBS) 10×
Reagent Final concentration Quantity or Volume
Tris-HCl 1M pH 8 500 mL
NaCl powder 7 g
Milli-Q H2O Up to 1 L
Total 1 L
Note: Store at room temperature.
Tris-buffered saline supplemented with Tween (TBST) 1×
Reagent Final concentration Quantity or Volume
TBS 10× 1× 100 mL
Tween 20 1 mL
Milli-Q H2O Up to 1 L
Total 1 L
Note: Store at room temperature.
10% separating electrophoresis gel
Reagent Final concentration Quantity or Volume
40% acrylamide/bisacrylamide 10% 2.5 mL
Tris-HCl 1.5M pH 8.8 2.9 mL
10% APS 100 μL
TEMED 10 μL
Sterile Braun water 4.5 mL
Total 10 mL (1 gel)
Note: TEMED and APS should be added last to start gel polymerization. Add 500 μL of isopropanol on top of the gel to straighten it during polymerization. Use immediately.
4% stacking electrophoresis gel
Reagent Final concentration Quantity or Volume
40% acrylamide/bisacrylamide 4% 328 μL
Tris-HCl 0.5M pH 6.8 830 μL
10% APS 25 μL
TEMED 3.8 µL
Milli-Q H2O 2.1 mL
Total 3.3 mL (1 gel)
Note: Remove the isopropanol covering the 10% gel. TEMED and APS should be added last to start gel polymerization. Use immediately.
Western blot running buffer
Reagent Final concentration Quantity or Volume
Tris-glycine SDS 10× 1× 200 mL
Milli-Q H2O 1.8 L
Total 2 L
Note: Store at room temperature.
Western blot transfer buffer
Reagent Final concentration Quantity or Volume
Tris-glycine 10× 1× 200 mL
Methanol 15% 300 mL
Milli-Q H2O 1.5 L
Total 2 L
Note: Store at 4 °C.
Laboratory supplies
T25 cm3 cell culture flasks (Thermo Fisher Scientific, catalog number: 169900)
6-well plates (Thermo Fisher Scientific, catalog number: 10578911)
5 mL serological pipettes (Gilson, catalog number: F110126)
LUNA cell counting slides (Logos biosystems, catalog number: NC1765657)
1.5 mL sterile Eppendorf safe-lock tubes, microtube (Eppendorf, catalog number: 0030120086)
2 mL sterile Eppendorf safe-lock tubes, microtube (Eppendorf, catalog number: 0030120094)
5 mL sterile open-top thin wall ultra-clear tubes (Beckman Coulter, catalog number: C14279)
50 mL Falcon tubes (Thermo Fisher Scientific, catalog number: 10203001)
Filtered 1,000 μL pipette tips (Thermo Fisher Scientific, catalog number: 11749855)
Filtered 200 μL pipette tips (Thermo Fisher Scientific, catalog number: 11782584)
Filtered 10 μL pipette tips (Thermo Fisher Scientific, catalog number: 94052000)
Gene Pulser/MicroPulser electroporation cuvettes, 0.4 cm gap (Bio-Rad, catalog number: 1652081)
Coated glass bottom fluorodishes (WPI, catalog number: FD35-100)
Polyvinylidene difluoride membranes (Thermo Fisher Scientific, catalog number: 88518)
Electrophoresis gel wrap glass plates set (CBS Scientific, catalog number: MGP100R)
Whatman 3 mm paper for western blot (Cytiva, catalog number: 3630-917)
Equipment
Biosafety cabinet Herasafe KSP Class II (Thermo Fisher Scientific, model: 1300 series A2)
Incubator set to 5% CO2 and 37 °C (Binder, series BD)
ThermoMixer compact (Eppendorf, model: F1.5)
LUNA-II brightfield cell counter (Logos Biosystems, model: C100-Pro)
Eppendorf centrifuge with a fixed-angle rotor for 2 mL Eppendorfs (Thermo Fisher Scientific, model: 5430G)
Optima L-80 XP ultracentrifuge (Beckman Coulter, model: 392051)
FinnPip adjustable volume (P10, P200, P1000) pipettes (Thermo Fisher Scientific, model: monocanal Finnpip)
Pipette controller (Brandtech, model: accu-jet S)
Gene Pulser Xcell electroporation system (Bio-Rad, model: 1652660)
STED super-resolution microscope (Abberior Instruments, model: Expert Line GmbH)
ChemiDoc touch imaging system (Bio-Rad, model: 1708370)
Electrophoresis generator (Apelex, model: PS304 XL)
Double-wide mini-blotter (CBS Scientific, model: EBU-402)
Software and datasets
Image acquisition and analysis
Imspector Image Acquisition & Analysis Software (Abberior, v16.3, 2023)
ImageJ (Fiji, v1.54h, 15/12/2023)
Spectragryph (v1.2.16, 14/07/2022)
GraphPad Prism (v8.3.0, 28/10/2019)
Figure conception and formatting
Inkscape (v1.3.2, 26/11/2023)
Biorender (https://app.biorender.com/)
Procedure
Cell culture and viral biofilm isolation
Cell culture
Note: HTLV-1 chronically infected T cells are cultivated in suspension in a vertical flask (see Figure 1A). The cell suspension (stock culture) is regularly amplified and stored at -150 °C in cryotubes containing fetal calf serum (FCS) supplemented with 10% DMSO. Cells need to be thawed and put in culture at least one week prior to the experiment.
Prepare complete RPMI medium: RPMI 10% FCS, 1% Penicillin/Streptomycin (P/S).
Warm complete RPMI medium and PBS at 37 °C using a water bath.
Pellet HTLV-1 chronically infected T cells from a stock culture at 1,000× g for 5 min.
Resuspend the cells in 10 mL of PBS in a 50 mL falcon tube (wash).
Spin the cell suspension at 1,000× g for 5 min.
Resuspend cells at a concentration of 0.5 × 106 cells/mL in 4 mL of complete RPMI medium.
Transfer the homogenized suspension into a T25 cm3 flask.
Incubate the flask vertically at 37 °C and 5% CO2 for 96 h (Figure 1A).
Note: The doubling time of the C91-PL cell line is 3 days. After 4 days of incubation at high cellular density (0.5 × 106 cells/mL), the cell count reaches approximately 1.25 million cells in 1 mL and therefore a total of 5 million cells in 4 mL.
Figure 1. Isolation of HTLV-1 biofilms from chronically infected T cells. A. Protocol for isolation of viral biofilms from chronically infected T cells (C91-PL). C91-PL cells were maintained in culture for 96 h to allow natural biofilm detachment in the supernatant. Step 1: Cells and large cellular debris were removed from the suspension by centrifugation at 1,000× g. Step 2: Biofilm isolation proceeded at 10,000× g. Step 3: Centrifugation at 100,000× g did not pellet more biofilms. B. Immunoblot probed for HTLV-1 Gagp19 showing Gag expression in viral biofilms isolated at 10,000× g (2) or 100,000× g (3).
Biofilm isolation using sequential centrifugations
Note: The centrifugation speeds that were tested (10,000× g and 100,000× g) to pellet viral biofilms are described below.
After 96 h of incubation, collect and split the cell suspension (without mixing) in two Eppendorf tubes of 2 mL.
Spin at 1,000× g for 5 min at 4 °C to pellet the cells and clarify the supernatant (Figure 1A.1).
Collect the supernatants (1) and transfer them into two new Eppendorf tubes of 2 mL.
Spin the supernatants (1) at 10,000× g for 40 min at 4 °C (Figure 1A.2).
Collect the supernatants (2) and transfer them into two open-top thin wall ultra-clear tubes of 5 mL.
In parallel, keep the 10,000× g pellets (2) for subsequent analyses.
For western blot, resuspend the pellets in a total volume of 60 μL of PBS with 0.2% Triton.
Incubate for 10 min at room temperature (RT) and store at 4 °C.
Spin the supernatants (2) at 100,000× g for 90 min at 4 °C (Figure 1A.3).
Discard the supernatants and keep the 100,000× g pellets (3) for subsequent analyses.
For the western blot, identical pellets are pooled and resuspended in a total volume of 60 μL of PBS with 0.2% Triton.
Incubate for 10 min at RT and store at 4 °C.
Keep samples at 4 °C for short-term storage (1 week) or -20 °C for long-term storage (several months).
Detection of isolated biofilms using western blot
Note: Prepare a 10% acrylamide/bisacrylamide gel topped by a 4% stacking gel (see Recipes).
All membrane incubation steps are performed under gentle agitation on a lab rocker.
Prepare each sample by mixing in new 1.5 mL Eppendorf tubes:
5 μL of biofilm suspension (10,000× g or 100,000× g fractions)
4 μL of Laemmli SDS 4× buffer
7 μL of Milli-Q H2O
Boil samples at 95 °C for 5 min.
Spin samples at 500× g for 1 min.
Load 4 μL of the prestained protein ladder into the first well of the electrophoresis gel.
Load 16 μL of each sample into the following wells.
Run migration at 80 V until the samples reach the stacking gel.
Run migration at 100 V until the samples reach the end of the running gel.
Pre-wet a PVDF membrane in methanol for 10 min (activation) and briefly rinse in transfer buffer.
Transfer the separated proteins onto the activated PVDF membrane using the wet transfer method at 400 mA for 1 h.
Incubate the membrane in 10 mL of TBST buffer supplemented with 5% milk for 30 min (membrane saturation).
Transfer the membrane in the primary antibody mix containing:
3 mL of 5% milk in TBS-T buffer
3 μL (1:1000) anti-HTLV-1 Gagp19 primary mouse antibody
Incubate overnight at 4 °C under gentle agitation.
Wash the membrane three times in 5% milk in TBS-T buffer for 10 min under gentle agitation.
Transfer the membrane in the secondary antibody mix containing:
10 mL of 5% milk in TBS-T buffer
3.3 μL (1:3,000) anti-mouse HRP-conjugated secondary antibody
Incubate at RT for 2 h under gentle agitation.
Wash the membrane three times in TBST buffer for 10 min under gentle agitation.
Reveal the membrane using the ECL prime kit (Amersham) following the manufacturer’s instructions.
Image the revealed membrane (Figure 1B) with the ChemiDoc imaging system using the following parameters:
Select Chemiluminescence mode.
Set the exposure time manually to 1s.
Note: If your signal is too weak to be detected with 1 s of exposure, select “Rapid Auto-exposure.”
Data analysis (Western blot): HTLV-1 Gagp19 proteins were detected in pellets obtained at 10,000× g (Figure 1B.2) by western blot, demonstrating that viral material has successfully been isolated with this speed. Gagp19 proteins were not detected at 100,000× g by western blot, suggesting that most viral biofilms were pelleted at centrifugation speeds ≤ 10,000× g. Therefore, 10,000× g speed was used to pellet viral biofilms for the following analyses (super-resolution microscopy).
Analysis of isolated biofilms using super-resolution STED microscopy
Note: To confirm the successful isolation of HTLV-1 biofilms, we chose to analyze the viral material pelleted at 10,000× g using stimulated emission-depletion (STED) fluorescence microscopy. This technique allows us to go below 100 nm in x, y resolution and therefore decipher the spatial organization of individual viral particles, measure their diameter, and check if they successfully incorporated viral markers (i.e., Env glycoproteins) (Figure 3).
Electroporation of the pGag-YFP plasmid into HTLV-1 chronically infected T cells
To visualize viral particles, we electroporated HTLV-1 chronically infected T cells with a DNA construct encoding the fluorescent HTLV-1 Gag-YFP protein, which is incorporated into the nascent virions (Figure 2A). This electroporation protocol was optimized for 8 × 106 cells mixed with 10 μg of plasmids.
Warm Opti-MEM (1×) reduced serum medium and PBS at 37 °C using a water bath.
Pellet HTLV-1 chronically infected T cells from a stock culture at 1,000× g for 5 min.
Resuspend cells in 10 mL of PBS in a 50 mL Falcon tube.
Spin the cell suspension at 1,000× g for 5 min (wash).
Count and resuspend cells at a concentration of 2 × 107 cells/mL in 1 mL of Opti-MEM medium.
Transfer 400 μL of this cell suspension (= 8 × 106 cells) into a 1.5 mL Eppendorf tube.
Add 10 μg of pHTLV-1 Gag-YFP plasmids.
Mix gently by pipetting up and down using a P1000 pipette.
Incubate at 37 °C for 30 min.
Mix again gently by pipetting up and down using a P1000 pipette.
Transfer the mix into a 0.4 cm electroporation cuvette.
Place the cuvette into the pod of the Gene Pulser Xcell electroporation system.
Use the following electroporation program:
Voltage: 180 V
Pulse length: 10 ms
Number of pulses: 3
Time between pulses: 1 s
Press the red Pulse button to start the program.
Wait until the electroporation (Figure 2A) is completed (a few seconds).
Transfer 200 μL of the cell suspension into 4 mL of prewarmed RPMI 10% FCS without P/S in a 10 mL Falcon tube.
Mix gently by pipetting up and down using a P1000 pipette.
Divide the cell suspension into two wells of a 6-well plate (volume per well = 2 mL).
Incubate at 37 °C and 5% CO2 for 96 h.
Note: The doubling time of the C91-PL cell line is 3 days. After 4 days of incubation at high cellular density (0.5 × 106 cells/mL), the cell count reaches approximately 1.25 million cells in 1 mL and therefore a total of 5 million cells in 4 mL. 60% of these cells die from the lack of nutrients, which allows spontaneous viral biofilm release.
Figure 2. Electroporation of HTLV-1 chronically infected T cells with a plasmid encoding HTLV-1 Gag-YFP fluorescent proteins. A. Scheme: Electroporation of HTLV-1 chronically infected T cells (C91-PL) with a plasmid encoding HTLV-1 Gag-YFP. Gag-YFP(+) biofilms isolated at 10,000× g are then plated on a poly-L-lysine-coated fluorodish for immunofluorescence staining and STED imaging (see below). B. Representative confocal microscopy images of living Gag-YFP(+) chronically infected T cells. Each image is a projection of 10 successive optical z-slices of 1 μm. These images show the successful accumulation of Gag-YFP(+) virions (yellow) at the cell surface 24 h post-electroporation. Scale bars = 10 μm.
Remark: We confirmed in Arone et al. [12] that Gag-YFP proteins were successfully incorporated into Gag WT(+) and Env WT(+) virions. This demonstrates that the YFP fluorescent signal reflects wild-type virion accumulation into HTLV-1 biofilms (DOI: 10.1128/mbio.01326-23).
Isolation and plating of Gag-YFP+ biofilms for super-resolution microscopy imaging
After 96 h of incubation, transfer each well into 2 mL Eppendorf tubes (without mixing).
Spin them at 1,000× g for 5 min at 4 °C to pellet the cells (Figure 1A.1) and clarify the supernatant.
Collect the supernatants and transfer them into two new Eppendorf tubes of 2 mL.
Spin the supernatants at 10,000× g for 10 min at 4 °C to pellet viral biofilms as in Figure 1A.2.
Discard the supernatants, resuspend, and pool pellets containing biofilms in a total volume of 60 μL of TNE buffer for storage.
Note: At this point, you can keep the samples at 4 °C for short-term storage (1 week) or -20 °C for long-term storage (several months).
Note: The percentage of cells that dissociates their biofilm is very challenging to estimate. On one hand, even if HTLV-1 chronically infected T cells are releasing biofilms, viral aggregates can still be detected at the cell surface (they are continuously producing viral particles/biofilms), so we cannot discriminate between cells that have “lost their biofilms” and cells that did not. On the other hand, not every cell carries a biofilm at a given timepoint, and these structures are variable in size.
Coat glass-bottom fluorodishes with poly-L-lysine using the following steps:
i. Add 500 μL of PBS containing 0.01% poly-L-lysine in the center of a fluorodish using a P1000 pipette.
ii. Incubate for 30 min at RT.
iii. Remove the poly-L-lysine coating solution.
iv. Wash the bottom of the fluorodish by gently adding and removing 500 μL of PBS.
v. Repeat the washing step three times.
vi. Remove all residual PBS.
Transfer 30 μL of the Gag-YFP(+) biofilms previously isolated into a new 1.5 mL Eppendorf.
Add 470 μL of TNE buffer and gently mix by pipetting up and down to dilute the biofilm suspension.
Transfer the mix (total volume = 500 μL) in the poly-L-lysine coated fluorodish (Figure 2).
Incubate for 30 min at RT in the dark to let the biofilms adhere to the coated glass.
Add 1.5 mL of TNE buffer and proceed to the fixation and staining steps (see below).
Staining of Gag-YFP+ biofilms for stimulated emission-depletion (STED) microscopy:
Note: STED microscopy principle: Using a depletion beam with a “donut-shaped” intensity profile, fluorophore emission can be selectively silenced in the periphery of an excited spot, thus effectively reducing the area of detected fluorescence, and improving the lateral resolution. This allows the detection of individual viral particles in biofilms.
Once biofilms have adhered to the coated glass (Figure 2), remove the TNE buffer.
Add 500 μL of PFA buffer (fixation).
Incubate for 15 min at RT.
Remove PFA and add 500 μL of freshly prepared 50 mM NH4Cl (quenching).
Incubate for 5 min at RT.
Remove NH4Cl and add 1 mL of PBS (wash).
Remove PBS and add 1 mL of PBS containing 0.05% saponin and 3% BSA (permeabilization/saturation buffer).
Incubate for 15 min at RT.
Remove permeabilization/saturation buffer and add the primary antibody mix containing:
i. 200 μL of permeabilization/saturation buffer
ii 1 μL (1:200) anti-YFP primary rabbit antibody
iii. 2 μL (1:100) anti-HTLV-1 Envgp46 primary mouse antibody
Incubate for 90 min at RT.
Remove the primary antibody mix.
Wash the sample by gently adding and removing 1 mL of permeabilization/saturation buffer.
Repeat the washing step three times.
Remove permeabilization/saturation buffer and add the secondary antibody mix containing:
200 μL of permeabilization/saturation buffer
2 μL (1:100) anti-rabbit IgG STAR 580 secondary antibody
2 μL (1:100) anti-mouse IgG STAR Red secondary antibody
Incubate for 90 min at RT in the dark.
Remove the secondary antibody mix.
Wash the sample by gently adding and removing 1 mL of PBS buffer.
Repeat the washing step three times.
Keep the sample in 2 mL of PBS at 4 °C in the dark until STED imaging (Figure 3A).
Figure 3. Imaging fluorescent HTLV-1 biofilms using super-resolution STED microscopy. A. Representative STED microscopy images of fixed Gag-YFP(+) biofilms pelleted at 10,000× g and stained for the viral envelope protein (Env). Scale bars = 500 nm. B. The white cross-section, shown in A, is plotted next to the corresponding image (left graph) where the three yellow peaks correspond to individual viral particles (FWHM = 148 nm, 150 nm, and 144 nm). The right plot shows the FWHM distribution of n = 25 cross-sections of Gag-YFP(+) particles and represents the distribution of particles’ diameter. FWHM: full width at half maximum of the Gag-YFP fluorescent signal peak, measured using Spectragryph software. SD: standard deviation of the mean.
Imaging parameters (STED): Dual-color STED 2D images (Figure 3B) were acquired on an STED microscope equipped with a 100× oil objective using 580 nm (Star-Orange) and 630 nm (Star-Red) excitation laser sources, coupled with a pulsed 775 nm STED laser. Using 25% of STED laser power, we could obtain a lateral resolution of less than 100 nm. All images were processed with ImageJ software. (DOI: 10.1128/mbio.01326-23.)
Image analysis (STED): Using STED microscopy, we were able to detect Gag-YFP(+) structures in the 10,000× g isolates (Figure 3A). As shown in the representative images in Figure 3A, Gag-YFP(+) signal corresponds to spherical individual particles with a mean diameter (calculated with the full width at half maximum, FWHM, of the fluorescent peak) of 174 nm ± 19 nm (Figure 3B). This size is consistent with the diameter of HTLV-1 virions measured by cryogenic transmission electron microscopy [15]. Moreover, Gag-YFP(+) viral particles successfully incorporated Env glycoproteins (Figure 3A): most Gag-YFP(+) viral cores were in close vicinity with Env signal, which was confirmed in Arone et al. [12]. Although some particles did not exhibit Env glycoproteins at their surface, it is known that HTLV-1 particles contain variable numbers of Env molecules, most of which are unevenly distributed in the viral lipidic envelope [16]. Overall, our results demonstrate the successful isolation of HTLV-1 biofilms containing unaltered Gag(+)/Env(+) viral particles. These isolated biofilms were also analyzed using atomic force microscopy and mass spectrometry by Arone et al. [12].
Validation of protocol
The protocol has been successfully employed in various studies in the lab, as well as in our most recent publication (Arone et al. [12], DOI: 10.1128/mbio.01326-23; Figure 1, Figure 2). In this study, HTLV-1 biofilms were isolated using this protocol and processed by mass spectrometry to do a large-scale identification of biofilm components.
General notes and troubleshooting
In step A.1(f), be sure to resuspend HTLV-1 chronically infected T cells (C91-PL) at a concentration of 0.5 × 106 cells/mL and to incubate the T25 cm3 flask vertically, as high cell density is required for them to survive and grow properly.
In step A.2(a), ensure that cell density/viability has not dropped before harvesting viral biofilms, as it could directly impact the quantity of biofilms collected.
In steps A.2(e, g), pelleted viral biofilms appear as thin white deposits on the bottom of the tubes: try to carefully remove supernatants using a P1000 pipette to avoid losing viral material. Also, remove any residual liquid before resuspending biofilms to avoid any excessive dilution.
In step C1(j), be sure to have a homogenous cell suspension when launching the electroporation step, as they tend to form clusters very rapidly (<10 min). Clusters reduce electroporation efficiency, as the surface area subjected to the electric field decreases when cells are not well dispersed.
In step C1(p), avoid pipetting cellular debris that accumulate at the surface of the suspension after electroporation by plunging the tip at the bottom of the cuvette.
In step C3, always avoid pipetting directly on biofilms during sample preparation for STED imaging, so they are not detached from the glass slides.
Acknowledgments
The authors would like to thank Marie-Pierre Blanchard (MRI, CNRS Montpellier, France) for STED microscopy training and Frederic Eghaian (Abberior, Germany) for providing all STED-compatible secondary antibodies. We also acknowledge the Montpellier Imaging Center for Microscopy (MRI) and the CEMIPAI BSL3 facility for providing excellent working conditions. The illustrations were created with BioRender.com ([email protected]). This work was supported by the French Agency for Research on AIDS and Viral Hepatitis (grant ANRS0016); institutional funds from the Centre National de la Recherche Scientifique (CNRS); and a 3-year CBS2 Ph.D. fellowship from Montpellier University (UM, France). This protocol was described and validated in the original research paper entitled “HTLV-1 biofilm polarization maintained by tetraspanin CD82 is required for efficient viral transmission” (Arone et al. [12], DOI: 10.1128/mbio.01326-23).
Competing interests
The authors declare no competing interests.
References
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Donegan, E., Lee, H., Operskalski, E., Shaw, G., Kleinman, S., Busch, M., Stevens, C., Schiff, E., Nowicki, M., Hollingsworth, C., et al. (1994). Transfusion transmission of retroviruses: human T‐lymphotropic virus types I and II compared with human immunodeficiency virus type 1. Transfusion (Paris). 34(6): 478–483. https://doi.org/10.1046/j.1537-2995.1994.34694295061.x
Miyamoto, K., Tomita, N., Ishii, A., Nishizaki, T., Kitajima, K., Tanaka, T., Nakamura, T., Watanabe, S. and Oda, T. (1984). Transformation of atla‐negative leukocytes by blood components from anti‐ATLA‐positive donors In vitro. Int J Cancer. 33(6): 721–725. https://doi.org/10.1002/ijc.2910330603
Pique, C. and Jones, K. S. (2012). Pathways of cell-cell transmission of HTLV-1. Front Microbiol. 3: e00378. https://doi.org/10.3389/fmicb.2012.00378
Maali, Y., Journo, C., Mahieux, R. and Dutartre, H. (2020). Microbial Biofilms: Human T-cell Leukemia Virus Type 1 First in Line for Viral Biofilm but Far Behind Bacterial Biofilms. Front Microbiol. 11: e02041. https://doi.org/10.3389/fmicb.2020.02041
Pais-Correia, A. M., Sachse, M., Guadagnini, S., Robbiati, V., Lasserre, R., Gessain, A., Gout, O., Alcover, A. and Thoulouze, M. I. (2009). Biofilm-like extracellular viral assemblies mediate HTLV-1 cell-to-cell transmission at virological synapses. Nat Med. 16(1): 83–89. https://doi.org/10.1038/nm.2065
Thoulouze, M. I. and Alcover, A. (2011). Can viruses form biofilms?. Trends Microbiol. 19(6): 257–262. https://doi.org/10.1016/j.tim.2011.03.002
Alais, S., Mahieux, R. and Dutartre, H. (2015). Viral Source-Independent High Susceptibility of Dendritic Cells to Human T-Cell Leukemia Virus Type 1 Infection Compared to That of T Lymphocytes. J Virol. 89(20): 10580–10590. https://doi.org/10.1128/jvi.01799-15
Arone, C., Martial, S., Burlaud-Gaillard, J., Thoulouze, M. I., Roingeard, P., Dutartre, H. and Muriaux, D. (2023). HTLV-1 biofilm polarization maintained by tetraspanin CD82 is required for efficient viral transmission. mBio. 14(6): e01326–23. https://doi.org/10.1128/mbio.01326-23
Arone, C., Dibsy, R., Inamdar, K., Lyonnais, S., Arhel, N. J., Favard, C. and Muriaux, D. (2021). Illuminating the nanoscopic world of viruses by fluorescence super-resolution microscopy. Virologie. 25(3): 47–60. https://doi.org/10.1684/vir.2021.0908
Heidecker, G., Lloyd, P. A., Fox, K., Nagashima, K. and Derse, D. (2004). Late Assembly Motifs of Human T-Cell Leukemia Virus Type 1 and Their Relative Roles in Particle Release. J Virol. 78(12): 6636–6648. https://doi.org/10.1128/jvi.78.12.6636-6648.2004
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Article Information
Publication history
Received: May 4, 2024
Accepted: Oct 10, 2024
Available online: Nov 18, 2024
Published: Dec 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Category
Immunology > Immune cell staining > Immunodetection
Cell Biology > Cell isolation and culture > Virus isolation
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5,153 | https://bio-protocol.org/en/bpdetail?id=5153&type=0 | # Bio-Protocol Content
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This is a correction notice. See the corrected protocol.
Peer-reviewed
Correction Notice: The on-Site Monitoring and Specimen-Making of Ectoparasites on Rodents and Other Small Mammals
PY Peng-Wu Yin
XG Xian-Guo Guo
WS Wen-Yu Song
WD Wen-Ge Dong
YL Yan Lv
DJ Dao-Chao Jin
Published: Nov 20, 2024
DOI: 10.21769/BioProtoc.5153 Views: 84
Reviewed by: Zeeshan Banday Anonymous reviewer(s)
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The following post-publication correction has been made to “The on-Site Monitoring and Specimen-Making of Ectoparasites on Rodents and Other Small Mammals. Bio-protocol (2024) 14(21): e5104. DOI: 10.21769/BioProtoc.5104” (https://bio-protocol.org/e5104):
In Figure 2, the label “Posture specimen (Rattus tanezumi) (modified by Pan et al. [30])” has been corrected to “Posture Specimen (Rattus norvegicus).”
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Published: Nov 20, 2024
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© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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5,154 | https://bio-protocol.org/en/bpdetail?id=5154&type=0 | # Bio-Protocol Content
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A Highly Efficient System for Separating Glandular and Non-glandular Trichome of Cucumber Fruit for Transcriptomic and Metabolomic Analysis
LS Lei Sun *
ZF Zhongxuan Feng *
FW Fang Wang
YQ Yu Qi
MA Menghang An
LY Lin Yang
MF Min Feng
MW Mingqi Wang
HR Huazhong Ren
XL Xingwang Liu
(*contributed equally to this work)
Published: Vol 15, Iss 1, Jan 5, 2025
DOI: 10.21769/BioProtoc.5154 Views: 189
Reviewed by: Wenrong He Kailiang Bo Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Plant Physiology Aug 2023
Abstract
Cucumber (Cucumis sativus) trichomes play a critical role in resisting external biological and abiotic stresses. Glandular trichomes are particularly significant as they serve as sites for the synthesis and secretion of secondary metabolites, while non-glandular trichomes are pivotal for determining the appearance quality of cucumbers. However, current methods for separating trichomes encounter challenges such as low efficiency and insufficient accuracy, limiting their applicability in multi-omics sequencing studies. This protocol introduces an efficient system designed for the precise separation of glandular and non-glandular trichomes from cucumber fruit. The process begins with the pre-cooling of sorbitol buffer or ethanol solution and the RNA-free treatment of laboratory supplies, followed by sterilization and pre-cooling. After filling glass bottles with pre-cooling buffer and glass beads, cucumber ovaries are then placed in the glass bottles and the trichome is harvested by bead-beating method. The separation process involves sequential filtration through various steel sieves and centrifugation to separate trichomes. The separated trichomes obtained from this method are well-suited for subsequent multi-omics sequencing analyses. This protocol achieved high precision in separating glandular and non-glandular trichomes, significantly enhancing the efficiency of separation and sample collection processes. This advancement not only addresses existing limitations but also facilitates comprehensive studies aimed at exploring the genetic and biochemical diversity present within cucumber trichomes, thereby opening avenues for broader agricultural and biological research applications.
Key features
• Use cucumber fruits on the day of flowering.
• Pre-cooling and RNA-free treatment ensure supply quality and purity.
• Efficiently separate glandular and non-glandular trichomes.
• Trichome samples are suitable for multi-omics sequencing analysis.
Keywords: Cucumber fruit Glandular trichome Non-glandular trichome Precise separation Bead beating Trichome harvest
Graphical overview
Background
With the advancement of biotechnology, there has been a dramatic increase in the number of methods used to separate trichomes in order to study their development, biosynthesis, and metabolism within glandular trichomes. Over the years, methods of separating and collecting trichomes have made great progress in the research process [1].
Earlier methods, such as direct tweezing of trichome cells in tobacco (Nicotiana tabacum) [2] and tweezing of individual trichome cells in Arabidopsis thaliana after quick-freezing plant materials with liquid nitrogen [3] proved inefficient and inaccurate. In tomatoes (Solanum lycopersicum), Pasteur pipettes were employed to collect type VI glandular trichomes from stems and leaves [4]. Advances continued with innovations in mint (Mentha spp.), where chemical and physical methods, aided by buffer solutions, enhanced both plant material protection and glandular trichome yield [5]. These techniques have since been adapted and widely adopted across various plant species [6].
Recent years have seen the bead-beating method emerge as a predominant approach for tomato glandular trichome separation in buffered solutions, followed by filtration and density gradient centrifugation to distinguish between different glandular trichome types [7]. Similarly, this method has proven effective for enriching and purifying cannabis (Cannabis sativa) glandular trichomes [8]. Meanwhile, laser microdissection enables the direct separation of individual glandular trichomes, although its use is time-consuming, inefficient, expensive to collect large numbers of trichomes, and not suitable for large sampling requirements [9].
Previous methods, while foundational, have often suffered from limitations in sampling accuracy and efficiency, primarily focusing on glandular trichome separation while neglecting non-glandular trichomes. In addressing these limitations, this protocol focuses on utilizing North China type cucumber fruits on the day of flowering as the primary research materials. Emphasis is placed on optimizing trichome collection and separation methods to ensure adequate sampling and precise separation, encompassing both glandular and non-glandular trichomes. A substantial yield of trichomes was obtained using the bead-beating method, followed by successful separation of glandular and non-glandular trichomes using steel sieves of varying pore sizes. Non-glandular trichome contents were fully extracted via tissue grinding. Standard samples were prepared for subsequent multi-omics sequencing.
This protocol significantly enhances trichome separation efficiency, achieves precise glandular and non-glandular trichome separation, and serves as a valuable reference for trichome separation in other crops and tissues with a certain firmness. This protocol not only refines current methodologies but also underscores the critical role of precise trichome separation in advancing our understanding of plant biology and biotechnology applications.
Materials and reagents
Biological materials
1. North China type cucumber fruits on the day of flowering (Figure 1)
Figure 1. North China type cucumber fruit on the day of flowering
Reagents
1. Sorbitol (Sigma-Aldrich, CAS number: 50-70-4)
2. Tris-HCl (Sigma-Aldrich, CAS number: 1185-53-1)
3. Sucrose (Sigma-Aldrich, CAS number: 57-50-1)
4. KCl (Sigma-Aldrich, CAS number: 7447-40-7)
5. MgCl2 (Sigma-Aldrich, CAS number: 7786-30-3)
6. Succinic acid (Sigma-Aldrich, CAS number: 110-15-6)
7. EGTA (Sigma-Aldrich, CAS number: 67-42-5)
8. K2HPO4 (Sigma-Aldrich, CAS number: 7758-11-4)
9. Triton X-100 (Sigma-Aldrich, CAS number: 9036-19-5)
10. Anhydrous ethanol (Sigma-Aldrich, CAS number: 64-17-5)
11. RNase remover I (Huayueyang Biotechnology, catalog number: 0416-100)
Solutions
1. Sorbitol buffer (see Recipes)
2. 70% (v/v) ethanol (see Recipes)
Recipes
1. Sorbitol buffer
Reagent Final concentration Quantity or Volume
Sorbitol 200 mM 18.217 g
Tris-HCl 50 mM 25 mL
Sucrose 20 mM 3.4229 g
KCl 10 mM 0.3725 g
MgCl2 5 mM 0.5083 g
Succinic acid 5 mM 0.2958 g
EGTA 1 mM 0.1902 g
K2HPO4 0.5 mM 0.0435 g
Triton X-100 0.015% (v/v) 0.075 mL
H2O n/a Up to 500 mL
Total n/a 500 mL
2. 70% (v/v) ethanol
Reagent Final concentration Quantity or Volume
Anhydrous ethanol 70% (v/v) 350 mL
H2O (RNA-free) n/a 150 mL
Total n/a 500 mL
Laboratory supplies
1. Reagent bottles (Fisher Scientific, catalog number: FB800500)
2. Glass beads (Sangon Biotech, catalog number: A500478)
3. 900 μm steel sieves (Shaoxing Shangyu Shengchao Instrument Equipment, catalog number: 20 Mu)
4. 60 μm steel sieves (Shaoxing Shangyu Shengchao Instrument Equipment, catalog number: 250 Mu)
5. 45 μm steel sieves (Shaoxing Shangyu Shengchao Instrument Equipment, catalog number: 325 Mu)
6. Tissue grinder (Sangon Biotech, catalog number: F519062)
7. 50 mL centrifuge tubes (Sangon Biotech, catalog number: F607788)
8. Petri dishes (Sigma-Aldrich, catalog number: BR455751)
Equipment
1. High-speed refrigerated centrifuge (Bioridge, model: TGL-18M)
2. Optical microscope (OLYMPUS, model: CX23)
Procedure
A. Buffer pre-cooling
1. If the samples are obtained for RNA extraction, prepare the 70% (v/v) ethanol solution in advance and pre-cool at -20 °C.
2. If the samples are obtained for metabolite determination, prepare the sorbitol buffer in advance and pre-cool at 4 °C.
B. Pretreatment of laboratory supplies
1. For RNA extraction, perform RNA-free treatment on all experimental supplies involved. Dilute RNase remover I with deionized water at a ratio of 1:1,000. Subsequently, immerse reagent bottles, glass beads, steel sieves, tissue grinder, 50 mL centrifuge tubes, and Petri dishes in this diluted solution for 30 min. Repeat this procedure once with a fresh RNase remover I dilution.
2. After immersing, wrap the supplies in tin foil and autoclave them at 121 °C for 15 min.
3. Dry the laboratory supplies and pre-cool at -20 °C.
C. Trichome harvest
1. Prepare glass bottles containing 30 g of glass beads and 250 mL of pre-cooling buffer. Samples are obtained for RNA extraction using 70% (v/v) ethanol and for metabolite determination using sorbitol solution.
2. On the day of flowering, harvest 25–30 developing ovaries from cucumber plants (Figure 2A). Cut the middle section to a length of approximately 1 cm (Figure 2B) and collect them in glass bottles.
Figure 2. Details of cutting and shaking operation. (A) Developing cucumber ovary on the day of flowering. Bar represents 1 cm. (B) Cucumber ovary after cutting. Bar represents 1 cm. (C) A cut cucumber ovary before shaking. (D) A cut cucumber ovary after shaking.
3. Shake the glass bottles vigorously for 10–15 min until the non-glandular trichomes are almost completely removed from the fruit peels, as visible to the naked eye (Figure 2C and D). Keep the samples at ice-cold temperatures.
D. Separating glandular and non-glandular trichome
1. Remove any remaining cucumber fruit tissues from the glass bottle.
2. Use a 900 μm steel sieve to filter the remaining solution (approximately 250 mL) from the glass bottle in step D1 into a Petri dish and collect the filtrate. Wash the glass bottle with 50 mL of the corresponding solution and collect the additional filtrate. The recovery volume of the filtrate should be higher than 95% of the volume of the added solution.
3. Use a 60 μm steel sieve to filter the filtrate (approximately 285 mL) obtained in step D2 into a new Petri dish and collect the filtrate. Wash the glass bottle with 50 mL of the corresponding solution and collect the additional filtrate. The recovery volume of the filtrate should be higher than 95% of the volume of the added solution.
4. Wash the residual materials on the 60 μm sieve with 50 mL of the corresponding solution and collect them.
5. Observe 20 μL of the recovered products from step D4 under an optical microscope and check that the collected products are non-glandular trichomes. Centrifuge the collected products (approximately 45 mL) in step D4 at 5,000× g for 5 min at 4 °C and remove all supernatants. The precipitate will consist of non-glandular trichomes (approximately 1.5 g).
6. Use a 45 μm steel sieve to filter the filtrate (approximately 320 mL) obtained in step D3 into a new Petri dish and collect the filtrate. Wash the glass bottle with 50 mL of the corresponding solution and collect the additional filtrate. The recovery volume of the filtrate should be higher than 95% of the volume of the added solution.
7. Observe 20 μL of the recovered products from step D6 under an optical microscope and check that the collected products are glandular trichomes. Centrifuge the filtrate (approximately 370 mL) collected in step D6 at 5,000× g for 5 min at 4 °C and remove all supernatants. The precipitate will consist of glandular trichomes (approximately 0.8 g) that are ready for subsequent use.
E. Grinding non-glandular trichome
1. Suspend the non-glandular trichome enrichment products in 800 μL of the corresponding solution and transfer to a tissue grinder. Keep the samples at ice-cold temperatures.
2. Wash the 50 mL centrifuge tube with 500 μL of the corresponding solution and transfer the wash to the tissue grinder. Keep the samples at ice-cold temperatures.
3. Grind the samples with the tissue grinder until no obvious particles remain, then transfer to a new 50 mL centrifuge tube. Keep the samples at ice-cold temperatures.
4. Centrifuge the ground product at 5,000× g for 5 min at 4 °C and remove the supernatant (the main component is the buffer used for extraction). The remaining precipitate consists of non-glandular trichomes ready for RNA extraction. Keep the samples at ice-cold temperatures.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article:
• Feng et al. [10]. Novel players in organogenesis and flavonoid biosynthesis in cucumber glandular trichomes. Plant Physiology (Figures 1–3).
In this paper, we successfully separated glandular and non-glandular trichomes of cucumber fruits. The concentration of total RNAs prepared from glandular and non-glandular trichome samples reached 900 and 60 ng/μL, respectively. We conducted subsequent glandular trichome metabolome sequencing and glandular and non-glandular trichome transcriptome sequencing and mined many genes related to the development and metabolism of cucumber trichomes. The above experimental results prove that our protocol is reliable.
General notes and troubleshooting
General notes
1. RNA sample preparation: Ensure all pretreatments are completed and maintain an ice-cold environment throughout the process.
2. Filtrate collection: The recovery rate of the filtrate at each step should be higher than 95% of the volume of the added solution.
3. Protocol scope: This protocol is designed for obtaining one set of samples. Adjust the sample preparation based on actual requirements.
4. Storage: If the collected products are not used immediately, store them at -80 °C.
Troubleshooting
Problem 1: After shaking for 15 min, many non-glandular trichomes remained on the surface of the fruit peels.
Possible cause: The number of glass beads is too small, or the diameter of glass beads is too small.
Solution: Increase the glass beads to more than 30 g or replace them with glass beads with a diameter of more than 1 mm.
Acknowledgments
This study was supported by the National Key Research and Development Program "Strategic Science and Technology Innovation Cooperation" Key Special Project (2023YFE0206900) and "The 2115 talent development program" of China Agricultural University. A special thanks to the original research paper by Feng et al. [10] (DOI: 10.1093/plphys/kiad236), which described and validated the protocol used in our study, for their invaluable contributions to the field.
Competing interests
There are no conflicts of interest or competing interests.
References
Feng, Z., Bartholomew, E. S., Liu, Z., Cui, Y., Dong, Y., Li, S., Wu, H., Ren, H. and Liu, X. (2021). Glandular trichomes: new focus on horticultural crops. Hortic Res. 8(1): 158.
Li, L., Zhao, Y., McCaig, B. C., Wingerd, B. A., Wang, J., Whalon, M. E., Pichersky, E. and Howe, G. A. (2004). The Tomato Homolog of CORONATINE-INSENSITIVE1 Is Required for the Maternal Control of Seed Maturation, Jasmonate-Signaled Defense Responses, and Glandular Trichome Development. Plant Cell. 16(1): 126–143.
Wienkoop, S., Zoeller, D., Ebert, B., Simon-Rosin, U., Fisahn, J., Glinski, M. and Weckwerth, W. (2004). Cell-specific protein profiling in Arabidopsis thaliana trichomes: identification of trichome-located proteins involved in sulfur metabolism and detoxification. Phytochemistry. 65(11): 1641–1649.
Xu, J., van Herwijnen, Z. O., Dräger, D. B., Sui, C., Haring, M. A. and Schuurink, R. C. (2018). SlMYC1 Regulates Type VI Glandular Trichome Formation and Terpene Biosynthesis in Tomato Glandular Cells. Plant Cell. 30(12): 2988–3005.
Gershenzon, J., McCaskill, D., Rajaonarivony, J. I., Mihaliak, C., Karp, F. and Croteau, R. (1992). Isolation of secretory cells from plant glandular trichomes and their use in biosynthetic studies of monoterpenes and other gland products. Anal Biochem. 200(1): 130–138.
Chen, L., Tian, N., Hu, M., Sandhu, D., Jin, Q., Gu, M., Zhang, X., Peng, Y., Zhang, J. and Chen, Z. (2022). Comparative transcriptome analysis reveals key pathways and genes involved in trichome development in tea plant (Camellia sinensis). Front Plant Sci. 13e997778.
Bergau, N., Bennewitz, S., Syrowatka, F., Hause, G. and Tissier, A. (2015). The development of type VI glandular trichomes in the cultivated tomato Solanum lycopersicum and a related wild species S. habrochaites. BMC Plant Biol. 15(1): 289.
Livingston, S. J., Quilichini, T. D., Booth, J. K., Wong, D. C. J., Rensing, K. H., J., Laflamme‐Yonkman, Castellarin, S. D., Bohlmann, J., Page, J. E. and Samuels, A. L. (2020). Cannabis glandular trichomes alter morphology and metabolite content during flower maturation. Plant J. 101(1): 37–56.
Happyana, N., Agnolet, S., Muntendam, R., Van Dam, A., Schneider, B. and Kayser, O. (2013). Analysis of cannabinoids in laser-microdissected trichomes of medicinal Cannabis sativa using LCMS and cryogenic NMR. Phytochemistry. 8751–59.
Feng, Z., Sun, L., Dong, M., Fan, S., Shi, K., Qu, Y., Zhu, L., Shi, J., Wang, W. and Liu, Y. (2023). Novel players in organogenesis and flavonoid biosynthesis in cucumber glandular trichomes. Plant Physiol. 192(4): 2723–2736.
Article Information
Publication history
Received: Jul 17, 2024
Accepted: Nov 3, 2024
Available online: Nov 20, 2024
Published: Jan 5, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
How to cite
Category
Plant Science > Plant biochemistry > RNA
Molecular Biology > RNA > RNA extraction
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Peer-reviewed
In Vitro Assay to Examine Osteoclast Resorptive Activity Under Estrogen Withdrawal
CF Cara Fiorino *
SO Safia Omer *
NG Nisha Gandhi
RH Rene E. Harrison
(*contributed equally to this work)
Published: Vol 15, Iss 1, Jan 5, 2025
DOI: 10.21769/BioProtoc.5155 Views: 219
Reviewed by: Athanas GuzhaWei DaiJaira Ferreira de Vasconcellos
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Original Research Article:
The authors used this protocol in International Journal of Molecular Sciences Jun 2024
Abstract
The bone is a highly dynamic organ that undergoes continuous remodeling through an intricate balance of bone formation and degradation. Hyperactivation of the bone-degrading cells, the osteoclasts (OCs), occurs in disease conditions and hormonal changes in females, resulting in osteoporosis, a disease characterized by altered microarchitecture of the bone tissue, and increased bone fragility. Thus, building robust assays to quantify OC resorptive activity to examine the molecular mechanisms underlying bone degradation is critical. Here, we establish an in vitro model to investigate the effect of estrogen withdrawal on OCs derived from the mouse macrophage RAW 264.7 cell line in a bone biomimetic microenvironment. This simple and robust model can also be adapted to examine the effect of drugs and genetic factors influencing OC resorptive activity in addition to being compatible with fluorescent imaging.
Key features
• A robust in vitro protocol that allows molecular and functional studies of mature osteoclasts in response to estrogen and its withdrawal.
• Generation of inorganic bone-mimetic substrates for culturing and examining osteoclast resorptive behavior.
• This quantitative image-based approach is compatible with brightfield and fluorescence microscopy to assess osteoclast resorptive activity.
Keywords: Estrogen withdrawal Biomimetic Osteoclast Differentiation Bone resorption Calcium phosphate coating Resorption pit analysis Microscopy-based quantification
Graphical overview
Background
Osteoclasts (OCs) are specialized, multinucleated cells responsible for the selective resorption of bone matrix [1,2]. In vivo osteoclast-driven resorption is followed by osteoblast (OB) matrix deposition, resulting in the continuous remodeling of bone [2,3]. The coordination between OC and OB activation relies on signaling from mechano-sensing osteocytes [2,3]. Disruption in this dynamic equilibrium produces disease states like osteoporosis [4,5]. Postmenopausal estrogen deficiency is a disrupting factor that shifts this equilibrium toward unregulated OC activity [3,6,7].
Estrogen (E2) deficiency de-regulates bone homeostasis; OB activity and lifespan are impaired, while increased osteoclast formation, resorptive activity, and lifespan contribute to net bone loss [3,7]. The effects of estrogen loss on OC formation and function have previously been studied using ovariectomized mouse models and bone marrow–derived osteoclasts in vitro [8,9]. Primary macrophages and OCs are challenging to work with and place limits on the biochemical and molecular analyses one can perform [10]. In vitro models of estrogen deficiency using OB and osteocyte cell lines exist [11–13]; however, this protocol establishes a simple and reproducible in vitro system to examine the effect of estrogen presence and withdrawal on OCs [14]. In contrast to existing models, using the murine RAW 264.7 cell line produces a large and consistent source of OCs, avoiding complicated extraction and propagation in favor of greater research output. This simple, robust method is a powerful tool to study osteoclast activation and function under postmenopausal conditions.
To measure OC activity, resorptive capacity can be assayed using coverslips coated with a thin layer of inorganic calcium phosphate. Maria et al. [15] thoroughly characterized and validated biomimetic calcium phosphate coating to study OC activity. Our protocol outlines the production of the coating and includes the culturing, staining, and subsequent imaging and analysis that can be performed. We provide this method as an alternative to using standard animal bone and dentine slices [16,17]. While suitable for resorption pit studies, bone and dentine slices do not allow for easy visualization of OCs remaining on the substrate [18]. In addition to quantifying OC resorptive activity, our lab has used biomimetic bone-coated coverslips combined with immunofluorescence to examine cytoskeletal elements, including F-actin ring morphology, the microtubule network, and centrosome de-clustering through inhibitor studies [14,19].
Materials and reagents
Biological materials
1. RAW 264.7 mouse macrophage cell line (American Type Culture Collection, catalog number: TIB-71)
Note: To maintain consistency across replicates, we recommend using RAW cells between passages 3 and 13. We have observed reduced OC formation in older passages.
Reagents
A. Bone mimetic production and staining reagents
1. Tris base (BioShop, catalog number: TRS001)
2. Hydrochloric acid solution, 1 N (HCl) (Thermo Fisher Scientific, catalog number: FLSA481)
3. Sodium hydroxide solution, 1 N (NaOH) (Thermo Fisher Scientific, catalog number: 124260010)
4. Calcium chloride dihydrate (CaCl2·2H2O) (BioShop, catalog number: CCL302)
5. Sodium chloride (NaCl) (BioShop, catalog number: CCL302)
6. Magnesium chloride hexahydrate (MgCl2·6H2O) (BioShop, catalog number: MAG510)
7. Disodium hydrogen phosphate dihydrate (Na2HPO4·2H2O) (MilliporeSigma, catalog number: 71643)
8. Sodium bicarbonate (NaHCO3) (BioShop, catalog number: SOB999)
9. Distilled water; provided by facility
10. Milli-Q water
11. 70% ethanol prepared from anhydrous or 95% ethyl alcohol
12. Sodium nitrate solution, 2.5% (AgNO3) (MilliporeSigma, catalog number: 85193)
13. Triton X-100 (BioShop, catalog number: TRX506)
B. Cell culture reagents
1. Dulbecco’s modified Eagle medium (DMEM) (Wisent Inc., catalog number: 319-005-CL)
2. Fetal bovine serum (FBS) (Wisent Inc., catalog number: 085-450)
3. Phosphate-buffered saline (PBS) (Wisent Inc., catalog number: 311-010-CL)
C. OC differentiation reagents
1. Alpha modified Eagle medium (AMEM) (Thermo Fisher Scientific, catalog number: 12571063)
2. 17β-estradiol (MilliporeSigma, catalog number: E8875)
3. Dimethyl sulfoxide (DMSO) (BioShop, catalog number: DMS555.250)
4. Recombinant receptor activator of nuclear factor kappa-B ligand (RANKL) was produced from BL21 Escherichia coli transformed with a pGEX-GST-hRANKL vector (a gift from Morris Manolson, Faculty of Dentistry, University of Toronto).
Commercial alternative: Mouse TRANCE recombinant protein (Thermo Fisher Scientific, catalog number: 315-11-10UG)
D. Osteoclast fixation and staining reagents
1. Paraformaldehyde (PFA) (Electron Microscopy, catalog number: 15710-S)
2. Permeabilization buffer (0.1% Triton X-100, 100 mM glycine in PBS) (BioShop, catalog number: GLN001)
3. Alexa Fluor 488 Phalloidin (Thermo Fisher Scientific, catalog number: 12379)
4. DAPI (Thermo Fisher Scientific, catalog number: D1306)
Solutions
Solutions required to prepare bone mimetic:
1. 50 mM Tris base stock solution, pH 7.4 (see Recipes)
2. Calcium stock solution (see Recipes)
3. Phosphate stock solution (see Recipes)
4. Calcium phosphate solution (see Recipes)
5. Simulated body fluid solution (SBF) (see Recipes)
6. Cell removal solution (see Recipes)
Recipes
1. Tris base stock solution (500 mL)
Reagent Final concentration Quantity or Volume
Tris base 50 mM 3.03 g
1 N HCl n/a see note*
Milli-Q water n/a see note*
Total n/a 500 mL
*Note: Add Tris base to 400 mL of Milli-Q water and 20 mL of 1 N HCl. Once dissolved, bring pH to 7.4 by slowly adding 0.05 mL increments of 1 N HCl. Bring the final buffer volume to 500 mL using Milli-Q water.
Store at room temperature for 1 month.
2. Calcium stock solution (100 mL)
Reagent Final concentration Quantity or Volume
CaCl2·2H2O 25 mM 0.37 g
NaCl 1.37 M 8.00 g
MgCl2·6H2O 15 mM 0.30 g
Tris base stock solution 50 mM see note*
Total n/a 100 mL
*Note: Add reagents to 90 mL of Tris base stock solution. Once dissolved, measure pH. Adjust to pH 7.38–7.4, if required, by slowly adding 0.05 mL increments of 1 N HCl or 1 N NaOH. Bring the final volume to 100 mL using Tris base stock solution.
Store at room temperature for 1 month.
3. Phosphate stock solution (100 mL)
Reagent Final concentration Quantity or Volume
Na2HPO4·2H2O 11.1 mM 0.20 g
NaHCO3 42 mM 0.35 g
Tris base stock solution 50 mM see note*
Total n/a 100 mL
*Note: Add reagents to 90 mL of Tris base stock solution. Once dissolved, measure pH. Adjust to pH 7.38–7.4, if required, by slowly adding 0.05 mL increments of 1 N HCl or 1 N NaOH. Bring the final volume to 100 mL using Tris base stock solution.
Store at room temperature for 1 month.
4. Calcium phosphate solution (250 mL)
Reagent Final concentration Quantity or Volume
Na2HPO4·2H2O 2.25 mM 0.10 g
CaCl2·2H2O 4 mM 0.15 g
NaCl 0.14 M 2.05 g
Tris base 50 mM 1.51 g
1 N HCl n/a see note*
Milli-Q water n/a see note*
Total n/a 250 mL
*Note: Add reagents to 200 mL of Milli-Q water and 10 mL of 1 N HCl. Once dissolved, bring pH to 7.4 by slowly adding 0.05 mL increments of 1 N HCl. Bring the final buffer volume to 250 mL.
Store at room temperature for 1 month.
5. Simulated body fluid solution (SBF)
Reagent Final concentration Quantity or Volume
Tris base stock solution n/a 25 mL
Calcium stock solution n/a 12.5 mL
Phosphate stock solution n/a 12.5 mL
Total n/a 50 mL; see note*
*Note: Make SBF immediately before adding to coverslips/wells. Maintain a ratio of 2:1:1 for Tris base:calcium:phosphate stock solutions and adjust the total volume as required for your experiment.
Make fresh and use immediately.
6. Cell removal solution
Reagent Final concentration Quantity or Volume
NaCl 1 M 5.84 g
Triton X-100 0.20 % 0.2 mL; see note*
Distilled water n/a 100 mL
Total n/a see note*
*Note: Prepare 1 M NaCl before adding the Triton X-100. Sterile-filtered 1 M NaCl can be prepared separately and stored at room temperature for 1 month.
Make fresh and use immediately.
Laboratory supplies
1. 12-well 18R glass coverslips (Electron Microscopy Sciences, catalog number: 72290-08)
2. 12-well tissue culture plates (Sarstedt, catalog number: 83.3921)
3. Parafilm (MilliporeSigma, catalog number: P7668)
4. 0.22 μm sterile bottle top filter (Sarstedt, catalog number: 83.3941.511)
5. T-75 cell culture flasks (Sarstedt, catalog number: 83.3911.002)
6. 5 and 10 mL disposable serological pipettes (Sarstedt, catalog number: 86.1253.001 and 86.1254.001, respectively)
7. 10, 200, 1,000 μL standard micropipette disposable tips
8. 1.5 mL standard microcentrifuge tubes
9. Cell scrapers (Sarstedt, catalog number: 83.3951)
10. 15 mL conical tubes (Sarstedt, catalog number: 62.554.205)
11. Microscope slides (Thermo Fisher Scientific, catalog number: 22037294)
12. DAKO (Agilent Technologies, catalog number: S302380-2)
13. Zeiss Immersion Oil 518F (Thermo Fisher Scientific, catalog number: 12-624-66A)
14. Slide folder (Thermo Fisher Scientific, catalog number: 12-587-10)
Equipment
1. Analytical balance (Mettler Toledo, catalog number: MS204S)
2. Magnetic stir plate (Thermo Fisher Scientific, catalog number: 1160049H)
3. 37 °C non-humidified incubator (Thermo Fisher Scientific, model: 637D)
4. 37 °C, 5% CO2 humidified incubator (Thermo Electron Corporation, model: HERACELL VIOS 160i)
5. 37 °C water bath (Thermo Fisher Scientific, model: ISOTEMP 110)
6. Vacuum aspirator (Integra-biosciences, catalog number: 158320)
7. Pipetman 2, 20, 200, and 1,000 μL (Gilson)
8. Autoclave; provided by research facility
9. Milli-Q Integral Water Purification System supplemented with Millipak® Express 0.22 μm membrane filter (Millipore, catalog number: MPGP04001)
10. pH meter (Thermo Fisher Scientific, model: AB150)
11. 500 and 1,000 mL glass beakers
12. 50 and 500 mL graduated cylinders
13. 250 and 500 mL glass bottles
14. Brightline hemacytometer (Electron Microscopy Sciences, catalog number: 100498-504)
15. Fine tip tweezers (Excelta, catalog number: 3C-SA-SE)
16. Inverted epifluorescence microscope with differential interference contrast (DIC) at 20× and 40× (Zeiss, model: AxioObserver Z1)
17. Inverted spinning disk confocal microscope with DIC at 40× 1.4 NA oil immersion lens (Quorum Technologies Inc., model: Quorum WaveFX-X1)
18. BSL-2 biosafety cabinet (Thermo Fisher Scientific, model: 1375 Series Class II)
Software and datasets
1. Zeiss Zen (3.1, blue edition, June 2012); license required
2. MetaMorph; license required
3. Fiji/ImageJ (2.3.0, May 2017); freely available
4. Prism GraphPad Software (7.03, March 2017); license required
Procedure
A. Preparation of a bone biomimetic microenvironment
Preparation of biomimetic bone substrate was first reported by Maria et al. [15] and Patntirapong et al. [20]. The inorganic bone layer produced using this protocol is chemically and structurally similar to in vivo bone mineral. This two-step method produces a stable, consistent crystal structure that is suitable for both OC differentiation and bone resorption [15,20]. Our method includes a maximum recommended shelf-life for the stock solutions and coated coverslips, in addition to a change in the preparation of the coverslips for cell culturing. A visual of the workflow involved is provided in Figure 1A.
Figure 1. Preparing biomimetic bone coverslips. A. This process involves the preparation of a biomimetic bone substrate following protocols adapted from Maria et al. [15] and Patntirapong et al. [20]. Key steps are (1) making stock solutions with precise pH requirements; (2) establishing a pre-calcification layer on coverslips using simulated body fluid (SBF); (3) producing the calcium phosphate top layer; and (4) sterilization and storage of coated coverslips for future experiments. B. Representative images of a growing nucleation layer (Day 2 and 3) and a completed calcium phosphate–coated coverslip. Images were acquired using DIC with a 20× objective. Scale bar = 100 μm.
1. Making stock solutions
Critical: The key to successfully preparing the final (SBF) solution is to ensure that all stock solutions have the correct pH. Adjust pH slowly and only use Milli-Q water when water is a required reagent.
Note: Choose an analytical balance with 100 mg minimum weight and 0.1 mg readability. A balance with a draft shield is recommended.
a. Autoclave three 250 mL bottles and one 500 mL bottle.
Note: Allow bottles to fully cool before adding solutions.
b. Prepare Tris base stock buffer solution in a 1,000 mL beaker using a magnetic stir rod and plate (see Recipe 1).
c. Keep Tris base stock solution in the beaker while preparing the calcium and phosphate stock solutions.
d. Prepare calcium stock solution using 90 mL of Tris base stock solution in a 500 mL beaker using a magnetic stir rod and plate (see Recipe 2).
e. Set aside the calcium stock solution, leaving it in the beaker.
f. Prepare phosphate stock solution in 90 mL of Tris base stock solution in a 500 mL beaker using a magnetic stir rod and plate (see Recipe 3).
g. Set aside the phosphate stock solution, leaving it in the beaker.
h. Prepare the calcium phosphate solution in a 500 mL beaker using a magnetic stir rod and plate (see Recipe 4).
i. Transfer each solution into a labeled and pre-sterilized glass bottle using a 0.22 μm bottle top filter.
Note 1: Use a vacuum aspirator to aid filtration. Attach aspirator tubing to the tapered connector of the bottle top filter.
Note 2: Performing this step in a BSL-2 biosafety cabinet is recommended to ensure sterility.
j. All sterile solutions should be kept at room temperature. Use within 1 month.
2. Establishing a pre-calcification/nucleation layer on coverslips with simulated body fluid
a. Prepare 12-well tissue culture plates with 18 mm coverslips.
Optional: Coverslips can be pre-sterilized in 70% ethanol and washed with sterile distilled water before addition to the tissue culture wells. This step can be prepared in advance.
b. Prepare simulated body fluid (SBF) in a 50 or 100 mL graduated cylinder according to recipe 5.
c. Seal the graduated cylinder with parafilm and mix with gentle inversion. Mix five times.
Note: The solution should remain clear after mixing. If the mixed solution appears cloudy or has a visible precipitate, then recheck the pH of each solution.
d. Add 1.0 mL of SBF to each well.
Note 1: Add the solution slowly to the side of the well.
Note 2: To achieve an even biomimetic layer across the coverslip, avoid circular movements with the plate, as this can cause mineral accumulation at the center of the well. Instead, employ a gentle side-to-side or up-and-down motion to evenly distribute the minerals.
e. Leave the tissue culture plates undisturbed at room temperature for 24 h.
f. Aspirate the SBF solution from each well carefully.
Note: Remove the solution from the side of the well, paying attention to avoid disturbing the amorphous calcification layer.
g. Repeat steps A2b–f for two additional days. A representative image of the nucleation layer on Day 2 and 3 of the process is provided in Figure 1B.
3. Production of the calcium phosphate top layer on coverslips
a. Aspirate the SBF solution from each well carefully and then add 1.0 mL of calcium phosphate solution (see Recipe 4) to the side of each well.
b. Leave the tissue culture plates undisturbed at room temperature for 24 h.
c. Slowly aspirate the calcium phosphate solution from the side of each well. A representative image of the completed top layer of calcium phosphate is provided in Figure 1B.
4. Sterilization and storage of coated coverslips housed within tissue culture plates
Note: Biomimetic substrate production is complete, and the following steps prepare the substrate for cell culture. Perform all remaining steps in a BSL-2 biosafety cabinet to maintain sterility.
a. Add 1.0 mL of 70% ethanol to the side of each well for 20 min to sterilize the substrate.
b. Aspirate the 70% ethanol from the side of each well and allow any remaining ethanol to evaporate for 1 h.
c. Wash the substrate-coated coverslips twice with autoclaved distilled water under sterile conditions.
d. Transfer the tissue culture plates to a 37 °C non-humidified incubator for 24 h.
e. Tissue culture plates with substrate-coated coverslips can be used for experiments immediately or stored at room temperature for later use. For long-term storage, seal the sides of the tissue culture plates with thin strips of parafilm to prevent contamination and stack the plates in a sealable bag. Use within 3 months.
B. Osteoclast culturing and estrogen withdrawal treatment
A simplified visual showing the timeline of events for osteoclast formation and estrogen treatment is provided in Figure 2A.
Figure 2. Protocol for osteoclast formation and treatment of cells under 17β-estradiol (E2) and E2-withdrawal (E2-WD) conditions. A. On Day 1, RAW 264.7 cells are seeded on biomimetic calcium phosphate–coated coverslips in AMEM + 10% FBS under the following conditions: 50 ng/mL RANKL for control and 50 ng/mL RANKL + 10 nM of 17β-estradiol for E2 and E2-WD. For the E2 condition, 10 nM of 17β-estradiol is added daily from Days 1 to 4. For E2-WD, 10 nM of 17β-estradiol is added daily on Days 1–2. On Day 3, AMEM + 50 ng/mL RANKL is replaced for all three conditions. On Day 5, the experiment is terminated for all conditions. B. Representative DIC images of the cell population from each day of treatment. Scale bar = 100 μm.
1. Passaging RAW 264.7 cells
Note 1: Perform all steps under sterile conditions, in a BSL-2 biosafety cabinet.
Note 2: RAW 264.7 cells are cultured and differentiated using medium supplemented with heat-inactivated FBS.
a. Maintain RAW 264.7 cells in T-75 cell culture flasks in DMEM supplemented with 10% FBS at 37 °C and 5% CO2 in a humidified incubator.
Note: Passage RAW cells twice a week as the flask population reaches 80%–90% confluency.
b. Remove old medium and add 10 mL of fresh prewarmed DMEM + 10% FBS medium. Using a sterile cell scraper, gently harvest cells from the tissue culture flask and, using a 5 or 10 mL disposable serological pipette, break cell clumps through gentle pipetting.
c. Transfer 1 mL of harvested RAW cells into a fresh flask containing 9 mL of prewarmed DMEM + 10% FBS. Mix by gentle pipetting to avoid creating bubbles before returning the flask to a 37 °C, 5% CO2, humidified incubator.
2. Prepare substrate-coated coverslips for cell culturing
a. Under sterile conditions, prepare a stock solution of 50% heat-inactivated FBS with AMEM.
b. Add 1 mL/well to tissue culture plates containing substrate-coated coverslips.
c. Place tissue culture plates in a 37 °C, 5% CO2, humidified incubator for 18–24 h.
3. Cell plating density and osteoclast differentiation medium
a. Transfer 2 mL of cells from Step B1 to a 15 mL conical tube containing 8 mL of prewarmed AMEM +10% FBS. Mix cells through gentle pipetting. For cell counting, use a sterile pipette tip to transfer 10 μL of cell suspension to a hemocytometer. Using a light microscope, focus on the grid lines of the hemocytometer using a 10× objective.
b. For cell seeding in a 12-well plate, prepare three labeled 15 mL conical tubes each containing 6 × 104 cells/mL in prewarmed AMEM supplemented with 10% FBS. Tube 1 is for the control (untreated osteoclasts) group, Tube 2 is for the continuous 17β-estradiol (E2) group, and Tube 3 is for E2-withdrawal (E2-WD) group.
Note 1: The total volume for each tube depends on the number of wells needed for each experimental condition.
Note 2: The number of cells used in seeding will change with the size of the well. For 24-well plates, we recommend using 10,000–15,000 cells/mL.
Note 3: To maintain consistency across replicates, using RAW cells between passage 3 and 13 is recommended. We have observed reduced osteoclast formation in older passages.
Note 4: Prepare three biological replicates for each condition.
c. For all three tubes, add RANKL to a final concentration of 50 ng/mL using fully thawed stock RANKL. Mix thoroughly with a 10 mL serological pipette.
Note: Maintain a master stock of 100 μg/mL RANKL in 50 μL aliquots at -20 °C. Thaw a single aliquot each week and store at 4 °C. Discard any remaining aliquot after 7 days. We find this avoids any potential challenges with RANKL degradation and loss of potency.
4. Estrogen withdrawal treatment
a. For the continuous E2 and E2-WD group, add 10 nM of thawed E2 to the previously prepared 15 mL conical tubes (step B3b), a concentration within the physiological range in mouse serum [21]. Mix thoroughly with a 10 mL pipette to evenly distribute the E2.
Note: E2 is dissolved in DMSO as a stock solution of 100 mM, divided into 100 μL aliquots, and stored at -20 °C. Use the resuspended E2 stock solution within 6 months. Before each experiment, prepare a working stock of 10 μM in DMSO. The working stock is used for the duration of a single experiment and is stored at -20 °C.
b. Remove tissue culture plates containing substrate-coated coverslips from the humidified incubator and aspirate the 50% heat-inactivated FBS with AMEM. Label tissue culture wells prior to the addition of appropriate cell suspension + treatment.
c. Add 1 mL of cell suspension + treatment/well (Day 1) and incubate the plates in a 37 °C, 5% CO2 humidified incubator.
d. For the E2 group, add 10 nM of E2 every day (Days 1–4). For cells in the E2-WD group, add 10 nM of E2 for Day 1 and 2.
Note: For all conditions, remove medium and replace with fresh AMEM supplemented with 10% FBS and 50 ng/mL of RANKL on Day 3.
e. On Day 3, for the E2-WD group, wash cells with 1 mL of AMEM three times to remove residual traces of E2. Add fresh AMEM + RANKL and continue to grow cells in the absence of E2 until Day 5. Representative DIC images of the OC population on each day of the experiment are provided in Figure 2B.
f. On Day 5, terminate the experiment by cell removal from the substrate (see below; section C) or by cell fixation, followed by permeabilization and fluorescent staining (see below; section D).
C. Osteoclast removal and calcium phosphate (modified Von Kossa) staining
1. Osteoclast removal from biomimetic bone substrate
a. Prepare cell removal solution according to Recipe 6.
b. Aspirate media from the 12-well plate.
c. Wash each well twice with 1.0 mL of distilled water.
d. Add 1 mL/well of cell removal solution. Leave for 5 min.
e. Aspirate the cell removal solution and wash each well with 1.0 mL of distilled water twice.
2. Calcium phosphate staining with silver nitrate (AgNO3) solution
a. In a fume hood, add 0.5 mL of 2.5% AgNO3 solution to each well.
Caution: AgNO3 solution is an eye and skin irritant. Perform all work in a fume hood.
b. Keep plates treated with 2.5% AgNO3 solution under direct light for 30 min to 1 h or until a dark-brown-to-black stain (see Figure 3A) has developed.
Figure 3. Steps to mount calcium phosphate-coated coverslip. A. Calcium phosphate–coated coverslip treated with 2.5% AgNO3 solution. Image shows a coated coverslip placed under direct light before (i) and after (ii) a 1 h incubation with 2.5% AgNO3 solution, developing a visible dark-brown-to-black stain. B. To mount the biomimetic-coated coverslip, apply a single drop of mounting medium to the glass slide surface before gently lowering the coated coverslip with the calcium phosphate–coated side up and away from the slide, then applying another drop of mounting medium and covering with a second clean coverslip.
c. In a fume hood, remove 2.5% AgNO3 solution with a pipette and dispose of it in a designated waste container.
Caution: AgNO3 is an acute and chronic aquatic hazard. Do not wash down drains. Identify the designated waste container with an Environmental Health and Safety–recognized chemical waste label and store the container according to directions from the facility’s Environmental Protection Services department.
d. Wash each well with 1.0 mL of distilled water twice. Pipette the wash volume into the designated waste container.
3. Imaging resorption pits
a. In a pop-up slide folder, place labeled glass slides and apply a small amount of DAKO mounting medium to the surface of the slide. Using fine-tip tweezers, place the edge of the coverslip over the mounting medium and carefully lower the coverslip to avoid creating bubbles.
Note: For improved resolution of the resorption pits, face the calcium phosphate–coated side away from the slide glass surface (Figure 3B). Apply a drop of mounting media on top of the calcium phosphate material and carefully mount another coverslip on top, so that the calcium phosphate–coated coverslip is sandwiched as depicted in Figure 3B. Let the assembly dry inside a closed folder for 24 h.
b. Using a 20× or 40× objective, image 5–10 sections of the coverslip with brightfield microscopy.
D. Osteoclast fixation and actin cytoskeleton staining
1. OC fixation and permeabilization
a. Aspirate media from the 12-well plate.
b. Wash each well twice with 1.0 mL of room-temperature PBS.
c. Add 0.5 mL/well of 4% PFA. Add the PFA to the side of the well. Leave the plate covered in darkness at room temperature for 20 min.
d. Aspirate the PFA from each well and wash each well twice with 1.0 mL of PBS.
Pause point: The plate can be sealed with parafilm and placed at 4 °C to continue staining at a later time. Store the plate at 4 °C with 1.0 mL of PBS/well and stain cells within one week of fixation.
e. Add 0.5 mL of permeabilization buffer to each well. Leave the plate covered in darkness at room temperature for 15 min.
f. Wash each well twice with 1.0 mL of room-temperature PBS.
2. Blocking non-specific binding
a. In a 15 mL conical tube, prepare a solution of 5% FBS in PBS.
b. Aspirate the PBS from each well. Add 0.5 mL of 5% FBS block to each well and incubate for 1 h at room temperature. Cover the 12-well plate with a box to prevent direct light exposure.
3. Actin and nuclei staining
Note: Prepare and store Phalloidin and DAPI according to the manufacturer’s instructions.
a. In a 15 mL conical tube, prepare the staining buffer as a solution of 2% FBS in PBS. Calculate the total volume of staining buffer needed. For a 12-well plate, use 0.5 mL/well.
Note: Include an extra sample in the total volume calculation to accommodate for pipetting errors. For example, prepare 6.5 mL of staining buffer for 12 wells.
b. Add a 1:250–1:500 dilution of Phalloidin 488 to the staining buffer prepared in step D3a.
c. Add 0.5 mL/well of Phalloidin staining solution, cover the plate, and leave for 1 h at room temperature. Avoid direct light exposure.
d. Aspirate the staining buffer and wash each well twice with 1.0 mL of room-temperature PBS and once with 1.0 mL of double-distilled water.
e. In a 15 mL conical tube, prepare a 1:1,000 dilution of DAPI (final concentration of 1–2 μg/mL) in double-distilled water.
f. Add 1 mL/well of DAPI staining solution, cover the plate, and leave for 10 min at room temperature. Avoid direct light exposure.
g. Aspirate the DAPI staining solution and wash the wells twice with 1.0 mL of double-distilled water.
4. Imaging osteoclasts
a. In a pop-up slide folder, place labeled glass slides and mount each coverslip using the method shown in Figure 3B.
b. Using a 40×–63× objective, image 5–10 sections of the coverslip with a fluorescence microscope that contains excitation and emission filters suitable for imaging the fluorescent probes, DAPI and Phalloidin 488. Examples of osteoclasts stained on calcium phosphate–coated coverslips and imaged with a fluorescence microscope are provided in Figure 4.
Figure 4. Osteoclasts in control conditions, fixed and stained on calcium phosphate–coated coverslips. Representative confocal images of OCs on each day of treatment. Actin (Phalloidin) and nuclei (DAPI) are shown. Inset, the biomimetic substrate is shown with DIC. Day 2 and 3 populations are mostly mononuclear cells. Osteoclasts (cells with 3 or more nuclei) are observed on Day 4 and 5. Scale bar = 100 μm.
Data analysis
Resorption pit analysis
1. Upload the resorption pit image into Fiji (Figure 5A).
Note: To download Fiji, go to the website (https://imagej.net/software/fiji/downloads). Fiji package is supported in MAC, Windows, or Linux. To install Fiji, select the appropriate file compatible with your machine. Once the download is complete, extract the compressed Fiji file and select the Fiji icon. No additional plugins are required for this analysis. Fiji will recognize most file extensions of microscope imaging software. However, should any problems arise, images can first be converted to TIF using the microscope imaging software.
Figure 5. Resorption pit analysis using Fiji software. A. Brightfield image of resorption pits converted to 8-bit in Fiji. B. Representative snapshot of the Threshold window with appropriate details selected in Fiji. Threshold sliders are adjusted until the resorption pits appear bright red and there is minimal red visible in the image background. C. Brightfield image with thresholding applied. Scale bar for A and C = 50 μm. D. Representative snapshot of Analyze Particles window with appropriate details selected in Fiji. The smallest allowable pit size is set by adjusting the size value (x-Infinity). The range should capture smaller resorption pits while excluding small variations (gaps) that are present in the substrate. A coverslip without cells can be used to estimate the size of the gaps. E. Outline of resorption pits with 5-Infinity chosen as size threshold. F. Outline of resorption pits with 50-Infinity chosen as size threshold. Notice that 50-Infinity compared to 5-Infinity avoids mistaking small variations in the substrate for resorption pits in the final measurement.
2. Convert the image to 8-bit (Image → Type → 8-bit).
3. Set the scale for the image using a known distance (as in a scale bar previously applied with the microscope imaging software). This will allow the user to present collected data in calibrated units like square micrometers (μm2). Use the line selection tool to draw a line along the scale bar. Choose Analyze → Set Scale. Adjust the values for “known distance” and “unit of length” according to the scale bar. Check the “Global” box to use this scale on subsequent images in the analysis set.
Note: This step applies if images do not contain embedded metadata.
4. Set the lower and upper threshold values for grayscale images using Image → Adjust → Threshold (Figure 5B and C). Use the “Default” method for thresholding. Adjust the sliders until the pits are completely red and minimal red is visible in the image background. “Dark background” remains checked since the pits are lighter than the background.
5. Analyze the pits using Analyze → Analyze Particles (Figure 5D). Begin with “set size” as 50-Infinity. The lower value is adjusted according to the smallest allowable pit size determined by the experimenter. Once decided, the lower value is maintained across all images.
Note: Select “Bare outlines” under the “show” dropdown to display the detected objects for visual verification (Figure 5D). An example of 5-Infinity (Figure 5E) and 50-Infinity (Figure 5F) is provided for comparison. Select the “Display results” option to collect readings on individual objects.
6. A summary table provides the number of resorption pits, total pit area, average size of pits, and the percentage area covered by the resorption pits. Transfer relevant data to a Microsoft Excel sheet. Record values from 10–15 fields/images per experimental condition.
7. Use Prism to plot measurements from the summary table into column data format and view the data as a graph (Figure 6). Alternatively, use Microsoft Excel to plot data. Depending on the number of conditions and data distribution, apply appropriate statistical tests.
Figure 6. Quantification of osteoclasts under E2-WD condition shows enhanced mineral resorption compared to resorption by osteoclasts treated continuously with E2. Graphs displaying the frequency distribution of the average number of resorption pits per field of view. Data are presented as the mean ± SEM from three biological replicates. p-values were calculated using the one-way ANOVA test followed by Tukey’s multiple comparison test. The high prevalence of smaller-sized pits indicates that estrogen suppresses osteoclast resorptive activity. Upon withdrawal of E2, osteoclasts regain their resorptive activity, resulting in more pits with larger areas, reaching levels similar to control cells. These findings align with a model where E2 withdrawal enhances bone degradation, similar to observations in postmenopausal females.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Gandhi et al. [14]. In vitro cell culture model for osteoclast activation during estrogen withdrawal. International Journal of Molecular Sciences (Figures 1 and 2).
Philip et al. [19]. Terminally differentiated osteoclasts organize centrosomes into large clusters for microtubule nucleation and bone resorption. Molecular Biology of the Cell (Figure 1B; 2B; 5A and D; 7A; 8).
Maria et al. [15]. Reproducible quantification of osteoclastic activity: Characterization of a biomimetic calcium phosphate assay. Journal of Biomedical Materials Research B: Applied Biomaterials (Figures 2, 4, 5 and 6).
General notes and troubleshooting
General notes
1. Commercially available murine RANKL (Thermo Fisher Scientific, catalog number: 315-11C-10UG) can be substituted for RANKL produced in-lab. Use the commercial RANKL at a final concentration of 100 ng/mL.
2. Perform serial dilution experiments whenever a new lab-prepared RANKL batch is used to confirm the optimal concentration for osteoclast formation. We suggest testing a range of 25–100 ng/mL.
3. Biomimetic-coated coverslips can be used with primary cells. We have successfully transferred osteoclasts produced from murine bone marrow–derived macrophages (BMDMs) onto biomimetic substrate for immunostaining purposes [19].
4. Distilled water and tap water are not appropriate for the creation of stock solutions. Our lab has tried both with no success. Milli-Q water is a critical reagent for the production of the calcium phosphate substrate.
5. Fluorescent staining can also be accomplished through methanol fixation. Apply 100% ice-cold methanol to coverslips and transfer the plates to the -20 °C for 7 min. Perform washes with 4 °C PBS before proceeding with blocking and staining at room temperature, while avoiding direct light exposure.
Troubleshooting
Problem 1: Poor osteoclast formation.
Possible cause(s): Cell passage, RANKL potency, and inaccurate cell density.
Solution 1: Confirm that RAW cells are between passage 3 and 13.
Solution 2: Use a fresh aliquot of RANKL or perform a serial dilution with a different RANKL lot.
Solution 3: Consider seeding cells at higher densities. For example, seed RAWs at 60,000, 70,000, and 80,000 cells/well to account for differences in cell counting accuracy.
Problem 2: Substrate lifts during silver nitrate staining.
Possible cause(s): Inappropriate washing before the addition of cell removal solution and 2.5% AgNO3 solution. Inappropriate storage of coated coverslips.
Solution 1: Wash cells and substrate with distilled water. Do not use PBS. We found that PBS can react with the silver nitrate solution and cause the substrate to lift.
Solution 2: Ensure that the coated coverslips are sealed during protracted storage. Changes in humidity can negatively impact substrate stability.
Problem 3: High background after staining.
Possible cause(s): Off-target binding to biomimetic coating.
Solution 1: Try blocking the coated coverslips with 5% FBS overnight at 4 °C. Additionally, wash coverslips thoroughly after antibody incubation. Gently rocking the plate may improve washing.
Solution 2: Try 2%–5% bovine serum albumin (BSA) as a blocking agent instead of FBS.
Acknowledgments
Graphics were created with BioRender and Adobe® Illustrator and Photoshop software. We thank Anisa Mazraeh and Anahita Karimzadeh for assistance in generating Figure 4. The biomimetic bone coating protocol is adapted from the original research paper by Maria et al. [15]. The combined estrogen withdrawal and bone microenvironment protocol is described and validated in the original research paper by Gandhi et al. [14]. S.O. is a recipient of a University of Toronto Provosts Postdoctoral Award and a Canadian Institutes of Health Research (CIHR)-REDI award (ED6-190720). R.E.H. is supported by a grant from CIHR (PJT-166084).
Competing interests
The authors declare no competing interests.
References
Boyle, W. J., Simonet, W. S. and Lacey, D. L. (2003). Osteoclast differentiation and activation. Nature. 423(6937): 337–342.
Kenkre, J. and Bassett, J. (2018). The bone remodelling cycle. Ann Clin Biochem. 55(3): 308–327.
Manolagas, S. C., (2000). Birth and Death of Bone Cells: Basic Regulatory Mechanisms and Implications for the Pathogenesis and Treatment of Osteoporosis. Endocr Rev. 21(2): 115–137.
Garnero, P., Sornay-Rendu, E., Chapuy, M. C. and Delmas, P. D. (1996). Increased bone turnover in late postmenopausal women is a major determinant of osteoporosis. J Bone Miner Res. 11(3): 337–349.
Raisz, L. G., (2005). Pathogenesis of osteoporosis: concepts, conflicts, and prospects. J Clin Invest. 115(12): 3318–3325.
Matsuo, K. and Irie, N. (2008). Osteoclast–osteoblast communication. Arch Biochem Biophys. 473(2): 201–209.
Weitzmann, M. N., (2006). Estrogen deficiency and bone loss: an inflammatory tale. J Clin Invest. 116(5): 1186–1194.
Chen, F., OuYang, Y., Ye, T., Ni, B. and Chen, A. (2014). Estrogen Inhibits RANKL-Induced Osteoclastic Differentiation by Increasing the Expression of TRPV5 Channel. J Cell Biochem. 115(4): 651–658.
Shevde, N. K., Bendixen, A. C., Dienger, K. M. and Pike, J. W. (2000). Estrogens suppress RANK ligand-induced osteoclast differentiation via a stromal cell independent mechanism involving c-Jun repression. Proc Natl Acad Sci USA. 97(14): 7829–7834.
Cuetara, B. L., Crotti, T. N., O'Donoghue, A. J. and McHugh, K. P. (2006). Cloning and characterization of osteoclast precursors from the RAW264.7 cell line. In Vitro Cell Dev Biol Anim. 42(7): 182–188.
Brennan, M. A., Haugh, M. G., O'Brien, F. J. and McNamara, L. M. (2014). Estrogen withdrawal from osteoblasts and osteocytes causes increased mineralization and apoptosis. Horm Metab Res. 46(8): 537–545.
Geoghegan, I. P., Hoey, D. A. and McNamara, L. M. (2019). Estrogen deficiency impairs integrin alpha(v)beta(3)-mediated mechanosensation by osteocytes and alters osteoclastogenic paracrine signalling. Sci Rep. 9(1): 4654.
Geoghegan, I. P., McNamara, L. M. and Hoey, D. A. (2021). Estrogen withdrawal alters cytoskeletal and primary ciliary dynamics resulting in increased Hedgehog and osteoclastogenic paracrine signalling in osteocytes. Sci Rep. 11(1): 9272.
Gandhi, N., Omer, S. and Harrison, R. E. (2024). In Vitro Cell Culture Model for Osteoclast Activation during Estrogen Withdrawal. Int J Mol Sci. 25(11): 6134.
Maria, S. M., Prukner, C., Sheikh, Z., Mueller, F., Barralet, J. E. and Komarova, S. V. (2014). Reproducible quantification of osteoclastic activity: characterization of a biomimetic calcium phosphate assay. J Biomed Mater Res B Appl Biomater. 102(5): 903–912.
Everts, V., Korper, W., Jansen, D. C., Steinfort, J., Lammerse, I., Heera, S., Docherty, A. J. and Beertsen, W. (1999). Functional heterogeneity of osteoclasts: matrix metalloproteinases participate in osteoclastic resorption of calvarial bone but not in resorption of long bone. FASEB J. 13(10): 1219–1230.
Jones, S. J., Boyde, A. and Ali, N. N. (1984). The resorption of biological and non-biological substrates by cultured avian and mammalian osteoclasts. Anat Embryol(Berl). 170(3): 247–256.
Boyde, A. and Jones, S. J. (1991). Pitfalls in pit measurement. Calcif Tissue Int. 49(2): 65–70.
Philip, R., Fiorino, C. and Harrison, R. E. (2022). Terminally differentiated osteoclasts organize centrosomes into large clusters for microtubule nucleation and bone resorption. Mol Biol Cell. 33(8): ar68.
Patntirapong, S., Habibovic, P. and Hauschka, P. V. (2009). Effects of soluble cobalt and cobalt incorporated into calcium phosphate layers on osteoclast differentiation and activation. Biomaterials. 30(4): 548–555.
Haisenleder, D. J., Schoenfelder, A. H., Marcinko, E. S., Geddis, L. M. and Marshall, J. C. (2011). Estimation of estradiol in mouse serum samples: evaluation of commercial estradiol immunoassays. Endocrinology. 152(11): 4443–4447.
Article Information
Publication history
Received: Aug 21, 2024
Accepted: Nov 3, 2024
Available online: Nov 20, 2024
Published: Jan 5, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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5,156 | https://bio-protocol.org/en/bpdetail?id=5156&type=0 | # Bio-Protocol Content
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Peer-reviewed
An Open-source Python Tool for Traction Force Microscopy on Micropatterned Substrates
AR Artur Ruppel
VM Vladimir Misiak
MB Martial Balland
Published: Vol 15, Iss 1, Jan 5, 2025
DOI: 10.21769/BioProtoc.5156 Views: 251
Reviewed by: Marc-Antoine SaniDjamel Eddine Chafai Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in eLIFE Aug 2023
Abstract
Cell-generated forces play a critical role in driving and regulating complex biological processes, such as cell migration and division and cell and tissue morphogenesis in development and disease. Traction force microscopy (TFM) is an established technique developed in the field of mechanobiology used to quantify cellular forces exerted on soft substrates and internal mechanical tissue stresses. TFM measures cell-generated traction forces in 2D or 3D environments with varying mechanical and biochemical properties. This technique involves embedding fiducial markers in the substrate, imaging substrate deformations caused by the cells, and using mathematical models to infer forces. This protocol compiles procedures from various previously published studies and software packages and describes how to perform TFM on 2D micropatterned substrates. Although not the focus of this protocol, the methods and software packages shown here also allow to perform monolayer stress microscopy (MSM), a method to calculate internal mechanical stress within the cells by modeling them as a thin plate with linear and homogeneous material properties. TFM and MSM are non-invasive methods capable of yielding spatially and temporally resolved force and stress maps with high throughput. As such, they enable the generation of rich datasets, which can provide valuable insights into the roles of cell-generated forces in various physiological and pathological processes.
Key features
• TFM and MSM protocol for 2D micropatterned polyacrylamide substrates, from sample preparation over imaging to data analysis with provided code.
• Sample preparation method is based on Tseng et al. [1].
• TFM analysis is done with Python custom code and is optimized for batch analysis of movies.
• MSM analysis is done with pyTFM from Bauer et al. [2].
Keywords: Traction force microscopy Monolayer stress microscopy Micropatterns Mechanobiology Biophysics
Graphical overview
Background
This protocol combines micropatterning, traction force microscopy (TFM), and monolayer stress microscopy (MSM), well-established techniques in mechanobiology, each with a rich history and extensive literature. As such, everything described in this protocol has been used and published by many scientists in the past. However, there is a lack of a comprehensive protocol integrating all three, spanning from sample preparation to data analysis, which this work aims to address.
Micropatterning is a tool that allows the imposition of geometrical boundary conditions onto cells by depositing adhesive proteins onto a substrate in specific spots. This tool has led to significant discoveries, showing that geometrical parameters are important in regulating complex biological processes, such as apoptosis [3], differentiation [4], or multicellular organization [5]. To combine micropatterning with TFM, Wang et al. developed a PDMS stencil-based protocol [6]. Later, Tseng et al. developed a photolithography-based method for micropatterned soft substrates, which the protocol here is based on [1].
The foundational study for TFM was published in 1980 [7], pioneering the use of soft substrates to visualize cell-generated traction forces, where the authors placed fibroblasts on a thin silicone sheet, which then showed visible wrinkling caused by cell-generated traction forces. A major improvement of this method consisted of visualizing the deformation of the substrate through fiducial markers, enabling detailed measurements of deformation maps of the substrates. The first studies using these improvements showed the first estimation of traction force maps for keratocytes [8] and fibroblasts [9].
The calculation of the traction force maps from the deformation maps is mathematically and computationally complicated due to the ill-posed nature of the problem. A first solution was proposed by Dembo et al. [10] and then, in a much more computationally efficient way, by Butler et al. [11]. For a more detailed review of the mathematical and computational foundations of TFM, see [12].
TFM primarily examines cell–substrate interactions, lacking insights into cell–cell interactions in multicellular systems. While simple force balance arguments suffice for cell doublets [13,14], more complex systems necessitate advanced approaches. This led to the development of monolayer stress microscopy (MSM) by Tambe et al. [15], which models cell monolayers as thin plates with homogeneous, linear material properties. Detailed discussions on assumptions and limitations are available [16]. Various researchers have independently developed solutions based on the same physical formulation of the problem [2,17,18].
Most scientists who publish TFM and/or MSM data develop their own analysis code, which is usually available only upon request. This requires significant technical expertise, which slows down the wide adoption of this technique in labs with a stronger focus on biological questions. For TFM, a wide variety of freely available analysis code now exists in the literature, such as Cellogram for reference-free TFM [19], TFMLAB for 4D TFM [20], pyTFM [2], Han Lab’s TFM [21], or JEasyTFM and iTACS for standard 2D TFM [22, 23]. For MSM, pyTFM by Bauer et al. and iTACS by Nguyen et al. are freely available [2, 23].
In order to cater to our specific TFM analysis needs, we developed our own Python TFM code, which is available at https://github.com/ArturRuppel/batchTFM. It is completely open-source and optimized for batch analysis of movies. It relies on numerous open-source Python packages, such as numpy and scipy, and most notably, relies on pyTFM for the MSM calculations.
Materials and reagents
Biological materials
1. Cell type of interest, opto-MDCK cells in our case. Wildtype MDCK cells were kindly provided by Prof. Yasuyuki Fujita, University of Kyoto, and then genetically modified by Dr. Manasi Kelkar and Dr. Guillaume Charras
Reagents
1. Adhesion protein of interest (usually fibronectin; Sigma-Aldrich, catalog number: F1141)
2. Fluorescent protein of choice (usually fibrinogen conjugated with, e.g., Alexa 546; Thermo Fisher, catalog number: F13192)
3. Dulbecco’s modified Eagle medium (DMEM) (Thermo Fisher, catalog number: 11965092)
4. Fetal bovine serum (FBS) (Thermo Fisher, catalog number: A5256801)
5. Penicillin-streptomycin, 5,000 U/mL (pen/strep) (Thermo Fisher, catalog number: 15070063)
6. Trypsin 2.5% (Thermo Fisher, catalog number: 15090046)
7. Ethylenediaminetetraacetic acid (EDTA) 0.5M (Thermo Fisher, catalog number: AM9260G)
8. Sterile phosphate-buffered saline (PBS) (Sigma-Aldrich, catalog number: 806552)
9. 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) (Thermo Fisher, catalog number: A14777.30)
10. Sodium bicarbonate (Sigma-Aldrich, catalog number: PHR3591)
11. Poly(L-lysine)-g-poly(ethylene glycol) (PLL-g-PEG) [SuSoS, catalog name: PLL(20)-g[3.5]- PEG(5)]
12. 2% Bis-acrylamide solution (Sigma-Aldrich, catalog number: M1533)
13. 40% Acrylamide solution (Sigma-Aldrich, catalog number: A4058)
14. Tetramethylethylenediamine (TEMED) (Sigma-Aldrich, catalog number: T9281)
15. Ammonium persulfate (APS) (Sigma-Aldrich, catalog number: A3678)
16. FluoSpheres, carboxylate-modified (e.g., dark red; Thermo Fisher, catalog number: F8807)
17. Bind silane (Sigma-Aldrich, catalog number: M6514)
18. 99.8% Ethanol (Thermo Fisher, catalog number: 445730025)
19. 99.7% Acetic acid (Sigma-Aldrich, catalog number: 695092)
20. Isopropanol (Fisher, catalog number: BP2618-1)
21. 96% ethanol (Carlo Erba, catalog number: 308646)
Solutions
1. 100 mM sodium bicarbonate, prepared with MilliQ water
2. 10 mM HEPES buffer, pH 7.4, prepared with MilliQ water and pH adjusted with NaOH
3. 10% acetic acid, diluted with MilliQ water
4. Cell culture medium (see Recipes)
5. Trypsinization solution (see Recipes)
6. 0.1 g/L PLL-g-PEG solution (see Recipes)
7. Polyacrylamide premix (see Recipes)
8. Silanization solution (see Recipes)
Recipes
1. Cell culture medium
Prepare in a sterile hood and then store in the fridge.
Reagent Final concentration Volume
DMEM 89% 445 mL
FBS 10% 50 mL
Pen/strep 1% 5 mL
Total 500 mL
2. Trypsinization solution
Prepare several aliquots in a sterile hood and then store in the freezer. Aliquots in use can be stored in the fridge for a few weeks.
Reagent Final concentration Volume
Trypsin 2.5% 0.25% 1 mL
EDTA 0.5M 0.5mM 0.01 mL
MilliQ water 8.99 mL
Total 10 mL
3. 0.1 g/L PLL-g-PEG solution
Filter the solution through a 0.2 μm filter and store the aliquots in the freezer. Aliquots in use can be stored in the fridge and should be used within ~10 days.
Reagent Final concentration Volume
PLL-g-PEG 1 mg 0.1 g/L
HEPES buffer 10mM pH 7.4 10mM 10 mL
Total 10 mL
4. Polyacrylamide premix
The exact recipe depends on the desired rigidity of the gel and can be found in [23]. Here, we provide the recipe for 20 kPa gels, also taken from [23]. Prepare in the fume hood. Can be stored in the fridge for years.
Reagent Final concentration Volume
40% Acrylamide solution 8% 2 mL
2% Bis-acrylamide solution 0.264% 1.32 mL
MilliQ water 6.68 mL
Total 10 mL
5. Silanization solution
Prepare under the fume hood and discard after use in appropriate chemical waste containers.
Reagent Final concentration Volume
Bind silane 0.35% 35 μL
10% acetic acid 0.324% 325 μL
99.8% ethanol 9.64 mL
Total 10 mL
Laboratory supplies
1. Borosilicate coverslips (32 and 25 mm) (VWR, catalog number: 631-0162 and 631-0171)
2. Coverslip cell chamber (e.g., from Aireka Cells or custom-made in a workshop. Technical drawings are available upon request)
3. Nitrogen spray gun (VWR, catalog number: ENTE421-42-11)
4. Tweezers (VWR, catalog number: 232-1220)
5. Squirt bottles (VWR, catalog number: 89141-044)
6. Photomask (Toppan)
7. Dish soap (argos, catalog number: 4202)
8. Kimwipe (Sigma, catalog number: Z671584)
9. Scalpel (Sigma, catalog number: S2896 and S2646)
10. 0.2 μm filters (Clearline, catalog number: 146560)
11. 5 mL syringes (Dutscher, catalog number: 6267268)
12. pH meter or pH test strips (Dutscher, catalog number: 30266626)
13. 145 mm Petri dish (Cellstar, catalog number: 639160)
Equipment
1. Deep UV lamp (Jelight UVO-Cleaner, model: Model 42)
2. Fume hood (Waldner, model: mc6 TA 1200X900.900 Fume Extraction Hood)
3. Plasma cleaner with oxygen supply (Diener, model: ASP-ATTO-M1)
4. MilliQ water purification system (Milli-Q, model: IQ7000)
5. Nikon Ti2-E epifluorescence microscope with perfect focus system (PFS) (Nikon, model: Ti2-E with PFS)
Software and datasets
1. All code and a test data set have been deposited to GitHub: https://github.com/ArturRuppel/batchTFM (commit af8114a, May 21, 2024)
Procedure
The procedure described hereafter is illustrated in Figure 1.
Figure 1. Schematic illustrating the procedure of a traction force microscopy (TFM) experiment on micropatterned samples
A. Coverslip silanization
This treatment ensures that the polyacrylamide gel will stick to the glass. All steps should be performed under a fume hood since bind silane is volatile and toxic.
1. Place as many coverslips as you can fit into the lid of a 145 mm Petri dish (because the lid can fit more coverslips than the base).
2. Pour 10 mL of silanization solution into the lid and cover with a second lid of a 145 mm Petri dish to avoid evaporation.
3. Incubate for at least 5 min.
4. Either dispose of the silanization into appropriate chemical waste or reuse it for a second batch of coverslips (prepared in another lid).
5. Rinse once or twice with 96% ethanol.
6. Remove coverslips while still covered with ethanol; otherwise, they will stick to the plastic. Take coverslips out with a tweezer and put them on a paper towel.
7. Dry carefully with another paper towel. Do not let the coverslips dry by themselves, as that will leave traces that could potentially be problematic for imaging.
8. Store silanized coverslips in a Petri dish for up to several months.
B. Clean coverslips for micropatterning
1. Clean coverslips with isopropanol and Kimwipes or use a coverslip rack, a beaker filled with isopropanol, and an ultrasonic bath to clean several coverslips at a time.
2. Use a nitrogen spray gun to dry the coverslips and remove dust.
3. Activate/clean the coverslips and the photomask in a plasma cleaner at 0.4 mbar with air atmosphere for 5 min.
C. Passivate coverslips with PLL-g-PEG
This step renders the coverslip anti-adhesive to both protein and cells. Subsequent removal of this passivation layer in specific places through photolithography allows for the patterning of the coverslip.
1. Put a drop of PLL-g-PEG solution on parafilm.
2. Take each coverslip with tweezers and flip it on the droplet with the plasma-activated side facing the PLL-g-PEG solution.
3. Let incubate for 30 min.
4. At the end of the incubation, lift the coverslips carefully without scratching the coating.
5. Clean the coverslips with a squirt bottle of MilliQ water. The PLL-g-PEG side should be very hydrophobic, and the water should go down easily.
6. Dry coverslips completely with a nitrogen spray gun or let them dry on a Kimwipe.
D. Deep UV burning and protein coating
This step removes the passivation layers from the previous step in specific places through photolithography, which allows for the patterning of the coverslip.
1. Make sure the photomask has been properly cleaned by the previous user (see section E).
2. Heat up the UV lamp for at least 5 min to reach a stable light intensity.
3. Rest the mask on a horizontal surface with the chrome side facing up.
4. Put a little drop of MilliQ water on the region of interest on the mask. Flip the coverslip onto the drop of water, with the PLL-g-PEG side facing the water. Remove excess water with a Kimwipe while applying pressure with your thumb. Wear gloves to not leave any traces on the mask or the coverslip. As much water as possible should be removed, so that there is good contact between the coverslip and the photomask. If good contact is established, you should see refraction patterns when looking at reflecting light on the coverslip.
5. Put the photomask in the warmed UV lamp with the coverslip-containing side facing away from the UV source. Place the photomask on three Petri dishes to avoid having the mask resting on the coverslips. Expose to UV light for 5 min.
6. Prepare the protein coating solution by diluting fibronectin to 20 μg/mL in 100 mM sodium bicarbonate.
7. After 5 min of UV light exposure, pour MilliQ water to help detach the coverslips from the mask. You can use a scalpel or tweezers (ideally with Teflon tips). When detaching the coverslips from the photomask, be very careful not to damage the mask. The patterned part of the coverslip is now hydrophilic, and the patterns can be seen when pouring water over the coverslip.
8. Put a drop of protein solution on parafilm (42 μL for 25 mm coverslips) and put the functionalized side of the coverslip on the droplet. Let it incubate for 30 min.
9. Pour MilliQ water over the coverslips, detach from the parafilm, and then rinse the coverslips with a squirt bottle of MilliQ water.
E. Mask cleaning
1. Wash the photomask with water and soap. Rub the surface gently with gloves.
2. Rinse the mask with plenty of deionized water and dry it with a nitrogen spray gun.
3. Rest the mask on a Kimwipe and pour isopropanol on it. Use another Kimwipe to thoroughly rub the surface of the mask.
4. Rinse the mask with 96% ethanol and dry it thoroughly with a nitrogen spray gun.
5. Put the mask in a plasma cleaner and pull a vacuum. Flush the chamber with oxygen for 5 min.
6. Pull a vacuum again and stabilize the pressure at 0.4 mbar. Activate the plasma for 10 min.
F. Polymerization and transfer of micropatterns to polyacrylamide gel
These steps should be performed quickly as polymerization starts as soon as APS is added, and if it proceeds too far before the micropatterned coverslip is placed, the drop might not spread completely. Once the acrylamide solution starts polymerizing, it is not toxic anymore and can be removed from the fume hood.
1. Mix 200 μL of polyacrylamide premix with 0.5 μL of FluoSpheres and vortex vigorously.
2. Put the premix in a vacuum chamber for 10 min to remove bubbles.
3. Ultrasonicate for 3 min.
4. Prepare a pipette with 42 μL and another with 1 μL.
5. Place silanized coverslips (functionalized surface facing up) on the top of a large Petri dish covered with parafilm.
6. Add 1 μL of TEMED and then 1 μL of APS and vortex vigorously.
7. Place a drop of 42 μL of the polyacrylamide solution onto the silanized coverslips and carefully place the micropatterned coverslip on top, with the patterned side facing the polyacrylamide solution.
8. Let the solution polymerize for 30 min.
9. Use the remaining acrylamide in the vial to check if polymerization was successful.
10. Pour deionized water over the polyacrylamide gels and let them hydrate for a few minutes.
11. Carefully detach the micropatterned coverslip with a scalpel. Do not force them apart, go carefully around the edges with the scalpel and allow the water to enter through the opening.
12. Place the coverslip in a 32 mm Petri dish or 6-well plate and rinse a few times with PBS until use.
G. Cell seeding
This protocol was used for cell seeding of MDCK cells expressing an optogenetic construct that allows activation of RhoA with the use of light. The specific parameters, such as incubation time for cell detachment, need to be adapted to the cell type.
1. Preheat cell detachment solution, sterile PBS, and cell culture medium to 37 °C in a water bath.
2. Put the coverslip with TFM gel in a coverslip cell chamber and rinse a few times with sterile PBS.
3. Add 1 mL of cell culture medium, put a lid from a Petri dish, and preheat in the incubator.
4. Remove the cell culture medium from the cell culture flask and rinse cells once or twice with sterile PBS to remove dead cells and cell debris.
5. Add cell detachment solution to cells. For a T25 cell culture flask, add 1 mL. Incubate for roughly 10–20 min. Frequent pipetting and tapping of the flask against the workbench helps to accelerate the process. Check frequently with a microscope the progress of cell detachment.
6. Transfer the cells in cell detachment solution to a 15 mL centrifugation tube and add 4 mL of cell culture medium.
7. Centrifuge at 234× g for 3 min to concentrate cells at the bottom of the tube.
8. Remove supernatant and resuspend in 5 mL of fresh cell culture medium.
9. Dilute part of the cell suspension to have the desired number of cells per sample in 1 mL of medium. Put some of the remaining cells back into cell culture.
10. To determine the number of cells to seed per sample, estimate the number of patterns on your coverslip and multiply by the number of cells per pattern. In our case, we had approximately 30,000 patterns, and we wanted to have one cell per pattern, so we seeded approximately 30,000 cells per sample.
11. Seed 1 mL of the correctly diluted cell suspension per sample. Pipetting tends to create liquid flow in the dish, which tends to concentrate cells in the center. To combat this and to get a more homogeneous seeding, pipette slowly while going in circles around the edge of the coverslip.
12. Incubate for 16–28 h to allow cells to divide on the patterns and form doublets or for 4–12 h to study single cells.
H. Imaging
We did all our imaging on a Nikon Ti-E2 microscope with an Orca Flash 4.0 sCMOS camera (Hamamatsu), a temperature control system set at 37 °C, a humidifier, and a CO2 controller, but any standard epifluorescence microscope can be used for TFM experiments. We strongly recommend the use of a drift correction system when acquiring movies, since small changes in focus can already strongly impact force measurements. We used and recommend physical drift correction systems, such as the PFS of Nikon, because of their speed and ease of use. If no microscope with such a system is available, drift correction systems based on real-time image analysis can be used, such as the drift correction software included in iTACS by Nguyen et al. [23]. Spinning disks or classical confocal microscopes can also be used. In this case, we recommend acquiring a small z-stack of a few micrometers around the top plane of the gel and averaging the images. If only one plane is taken, very small drifts, which usually happen even with drift correction systems, lead to images where the FluoSphere changes in size while going in and out of the focal plane, which can have an impact on the displacement measurements.
For the camera, a small pixel size is preferable over high sensitivity. The FluoSpheres in the gel are very bright, so high sensitivity is not necessary, and the pixel size is proportional to the resolution of displacement measurement. For the same reason, high magnifications (e.g., 60×) should be used when high precision is required. In principle, with particle tracking velocimetry, FluoSpheres movements can be measured with subpixel precision, but in practice, such a small signal would be indistinguishable from noise. On the other hand, too high displacements can lead to measurement errors. To get good displacement measurements, the rigidity of the gel should be chosen such that typical displacements fall between around 0.5 and 15 bead diameters (Figure 2).
Figure 2. Illustration of the impact of displacement magnitude on measurement accuracy. The left column shows artificially generated displacement maps, which were used to deform an example bead image. The middle column shows the result of the displacement measurements between the deformed and original images. The right column shows the difference between the measured and true displacement. The top row contains data from small displacements, the middle row from medium, “ideal” displacements, and the bottom row from high displacements.
Data analysis
We process our data with custom-written Python code, which can be downloaded from https://github.com/ArturRuppel/batchTFM. Note that this is an updated version compared to the code that was used in our study that this protocol is based on [24]. The old version was written in MATLAB and is no longer maintained by the author. The most important change is the switch from Particle Image Velocimetry + Particle Tracking Velocimetry to optical flow for the gel deformation measurement. We have compared the two versions and found no significant difference (see protocol validation).
First, create one folder for each position called “position”+index (i.e., “position0” for the first position and “position1” for the second one, as seen in the “test_data” folder on https://github.com/ArturRuppel/batchTFM). These folders need to contain a stack of images of the fluorescent beads while cells are attached to the hydrogel and one image of the beads after the removal of the cells when the hydrogel is in the relaxed state. At least one and up to three stacks of images of the cells need to be added as well. These images do not impact the data analysis; the code only corrects for translation through image registration. This allows to couple precise cell morphology and/or protein localization measurements with force localization. In the end, the script produces a movie of the first stack with the forces overlaid.
Analysis parameters
All data analysis parameters are found in the file “main.py” (Figure 3). First, set them according to your experiment and then run the script in your Python IDE (e.g., spyder).
The “path” parameter is a string, which should contain the path to the folder “position + index”. The different “name” parameters should contain a string with the filename of your images. We usually call “AK” (After Kill) the image of the beads in the relaxed state, “BK” (Before Kill) the movies of the beads in the stressed state, and “fluo” the image stack of the cells.
The “no_stacks” parameter represents the number of positions you want to analyze. “finterval” represents the time interval between two frames in minutes, “pixelsize” represents the physical size of a pixel in the bead images in micrometers, and “downsamplerate” represents the factor by which the final displacement and force maps are downsized. This serves mainly to conserve disk space. If set at 1, the displacement and force maps will have the same size as the bead images.
Then, choose which part of the analysis you want to launch on your data by uncommenting lines calling said functions (Figure 4). If one or several parts of the analysis are already done, they can be skipped by adding a “#” in front of the corresponding function.
Figure 3. Screenshot extracted from “main.py” that shows the parameters to set before running the code
Figure 4. Screenshot extracted from “main.py” showing the part where the steps to perform can be selected or unselected. Depending on which functions are enabled, more parameters need to be set in the “parameters to set” part of “main.py” (Figure 3).
The “preprocess_images” function takes all images described before and returns images with an adjusted contrast. Additionally, it aligns all bead images to the bead image in the relaxed state to correct translational xy drift from the microscope stage. It also applies the same translations to the additional cell images. This ensures that all images are well aligned at all time points, which is crucial if measurements on the cell images need to be combined with the TFM data. The outputs of this function are the processed image files, stored in a folder called “preprocessed_images” in the folder of the corresponding position.
The parameters “lower_threshold” and “upper_threshold” represent the percentile values for the contrast adjustment. The four different numbers correspond to the four different image stacks that can be put in. The first one corresponds to the bead images and the subsequent one for the cell images. The threshold values are the same for all the beads images, both in the relaxed and stressed state, so make sure that they were all acquired with the same imaging parameters.
The parameter “sigma_smooth” describes the size of the Gaussian kernel in pixels with which the images are smoothened. The “radius” variable corresponds to the size of a ball used for the “rolling ball background subtraction” algorithm (see doc skimage: https://scikit-image.org/docs/stable/auto_examples/segmentation/plot_rolling_ball.html). Smoothening and background subtraction are usually not necessary if images have a decent contrast and are likely to influence the measurement results. To skip these, set the radius to 0 and the smoothening window to 1. These features can still be useful if one wishes to qualitatively analyze failed experiments, or if acquiring high-contrast images is not possible. This part of the code returns a processed version of the user’s images and stacks in a folder called “preprocessed images.”
The “measure_displacement” function takes the images of the preprocessed images of the beads and returns the displacement matrices of the soft gel along the x- and y-axis. The output files are called “d_x.npy” and “d_y.npy” and are saved in the TFM_data folder created automatically in the “position+index” folder. These displacement matrices are obtained from estimations of the optical flow between each frame of the stressed bead images and the relaxed bead image. This function uses “optical_flow_tvl1” [26–28], a variational method that allows to compute an estimation of the optical flow between the bead images and returns the estimated displacement of the soft gel upon stress applied by the cells. The matrices returned by this function will be the size of the images of the beads divided by the “downsamplerate” defined earlier. (For example, if the size of the bead images is 600 × 600 pixels and the downsamplerate is 4, then the displacement matrix will be 150 × 150 pixels.) Each element of these matrices represents a pixel of the displacement map, and each pixel is associated with an estimation of the displacement of the soft gel quantified in micrometers along the x-axis for “d_x.npy” or the y-axis for “d_y.npy.”
The “calculate_forces” function solves the inverse problem to compute an estimation of the traction forces that were applied by the cells on the soft gel to observe the estimated displacements measured by the preceding function. It uses the classical approach of transforming the displacement field into the Fourier space to solve the corresponding equations. This approach is called Fourier Transform Traction Cytometry (FTTC) and was initially proposed by Butler et al. [11]. Before launching this function, define the Young modulus (called “E”) and the Poisson ratio (called “nu”) of the hydrogel used for the experiment you want to analyze. For example, in the associated paper from Ruppel et al. [24], a gel of approximately 19.66 kPa was used, so E = 19660. This gel is made of polyacrylamide, which has a Poisson ratio of 0.5. The traction forces are calculated via the solution of a linear system and from under-sampled data with potential acquisition noise. To correct for this, the function uses a regularization scheme, which effectively smoothens the output in the disfavor of overfitting the force field to a noisy displacement field. The regularization parameter is called “alpha” in the code, and we usually determine it empirically by setting it as low as possible without getting too much noise in the force fields. See Sabass et al. for a more detailed discussion of regularization in TFM calculations [29]. This function returns two matrices of the traction forces along the x- and y-axis defined arbitrarily by the horizontal and vertical axes of the images. These matrices have the same dimensions as the displacement matrices described earlier. Each element of these matrices is a spatial region associated with a value of the traction forces. As we have two matrices and two axes along which the tractions are applied, we can reconstruct a traction vector and represent the amplitude and the orientation of the forces applied by the cells on the substrate in a 2D traction force map.
The function "make_TFM_movies" is used to create movies or images of the traction force maps and to generate displacement maps. These maps represent the displacements of the gel under the cells using vectors, where the length and color indicate the magnitude of the displacement, and the orientation shows the direction of the bead displacement. The function also allows the user to get a version of the traction force map assembled with a potential brightfield or fluorescent channel to allow them to have a first impression of the link between the mechanical readouts calculated with the TFM and the morphology of the cells.
The function “apply_MSM” is used to perform monolayer stress microscopy, which takes the TFM data computed earlier and a mask with the contour of the cell layer to compute the internal mechanical stresses of the cell layer. It uses the MSM functions developed by the Fabry Lab, which can be found in the free Python package pyTFM [2]. MSM models the cell layer as a thin plate with linear and homogeneous material properties and then solves the associated partial differential equations to obtain internal mechanical stresses from the traction force data. For a detailed discussion of this method, see Tambe et al. [15]. The function “apply_MSM” returns a matrix that contains the Cauchy stress tensor for each pixel and each frame of the initial traction force image. The main diagonals of this stress tensor are the normal stresses in x- and in y-direction, respectively, and the off-diagonals correspond to the shear stresses.
Similar to “make_TFM_movies,” the “make_MSM_movies” function allows the user to generate movies that represent spatially the evolution of the internal stresses of the system from the MSM data. The function generates a movie for the two normal stresses along the x- and y-axes and also a global representation of the average normal stress in the system.
Validation of protocol
We validated the accuracy of our TFM code by analyzing a synthetic dataset. For this, we used code from Blumberg et al. [30] to generate an example displacement and traction force field that looks similar to what we typically observed in our experiments. Then, we applied this deformation field to an example bead image from one of our experiments. Finally, we used both the no longer supported MATLAB code and the new “batchTFM” code to find the deformation map between these two images. The results are shown in Figure 5. The main difference between these two versions of the script is the method used to find these deformation maps. The MATLAB code uses a combination of particle image velocimetry (PIV) combined with particle tracking velocimetry (PTV) to find this deformation map, and batchTFM uses an optical flow algorithm to find the same deformation map. As seen in Figure 5, the deformation maps obtained from these two methods are virtually identical. Figure 5 also shows the traction force maps calculated from the deformation maps. The same Fourier transform traction cytometry (FTTC) algorithm was used in both cases. In both cases, the original traction force maps are reproduced faithfully, albeit with some background force level and an underestimation of the peak force value. This is unavoidable and a direct consequence of the ill-posed nature of the force reconstruction problem and the hence necessary regularization scheme described earlier. To illustrate the output of the algorithm, Figure 6 shows force maps comparing cell doublets with single cells on H-shaped micropatterns.
Figure 5. The simulated displacement field on the top left was used to deform an example image of beads and these two images were then used to test the two different displacement measurement methods, namely particle image velocimetry (PIV) combined with particle tracking velocimetry (PTV) compared with optical flow (OF). The bottom row shows the correct traction stress field for the simulated data on the left. Taken from [31]. Scale bar is 10 μm.
Figure 6. Example of traction force data, comparing an example and the average traction of cell doublets and single cells on H-shaped micropatterns. Taken from [24]. Scale bar is 10 μm.
The MSM part of the analysis, which is part of the pyTFM software, has been validated by the developers of the software in [2].
General notes and troubleshooting
General notes
Setting up TFM experiments can be challenging; specifically, the analysis and interpretation of the data can become very technical. Once everything is set up, however, it works very reliably and can be used routinely. Do not hesitate to reach out if you need help setting up your TFM experiments.
Troubleshooting
Problem 1: The polyacrylamide gel does not polymerize properly.
Possible cause: The APS is not active anymore. APS is unstable in water and the powder tends to absorb water from the air.
Solution: Use fresh APS.
Problem 2: The micropatterns do not transfer well to the gel.
Possible cause: This usually happens only with micropatterns that have a big surface area (approximately >10.000 µm2).
Solution: Unfortunately, there is no simple solution to the problem. For big patterns, we recommend adding a crosslinker to the gel or using microcontact printing instead. Please contact us for more details if you have this issue.
Problem 3: The beads cluster under the micropattern.
Possible cause: This happens almost always and is due to unspecific interactions between your protein of interest and the beads.
Solution: We do not think that this is a problem as long as there are still enough beads in the immediate vicinity of the pattern. Outside of the pattern, the gel does not deform because there are no cells, so it is not a problem if there are fewer beads there.
Problem 4: There are a lot of displacement vectors at the edges of the image but there are no cells.
Possible cause: This is due to spherical aberrations at the edges of the image. This problem is exacerbated with cameras that have a large field of view.
Solution: Place your cells in the center of the image and crop the borders.
Problem 5: The displacement field is completely nonsensical, e.g., there is a rotational field in the whole field of view.
Possible cause: When trypsinizing the cells to take the reference images, it is extremely important to move the sample as little as possible. Small, translational drift is compensated by the algorithm, but large displacements or rotational movements will lead to nonsensical results or to an error message. Sometimes, the data can still be saved through additional image registration and/or manual alignment.
Solution: Do not touch the sample when trypsinizing the cells.
Problem 6: The resulting displacement field is extremely noisy.
Possible cause: The background in the bead image was not removed properly.
Solution: Either the contrast of the bead images is too low, or the threshold value in the algorithm is too low. In the former case, the experiment should be redone with higher light intensity and/or exposure time; in the latter case, only the threshold value in the algorithm needs to be adjusted.
Problem 7: The resulting displacement field is extremely noisy.
Possible cause: The forces generated by the cells are very weak.
Solution: Some cell types exert very little to no force. These can be very challenging to measure. Using softer gels can help, but typically this will also increase the noise. In any case, the smaller the forces one tries to measure, the more important it is to get good contrast and resolution in the imaging data.
Acknowledgments
This protocol describes the experiments done in our original study published in eLife [24]. We acknowledge the use of GPT 3.5 for generating a first draft of the Procedure section. We acknowledge the Agence Nationale de la Recherche (ANR-17-CE30-0032-01) for funding. Most importantly, we would like to thank Andreas Bauer and Ben Fabry for publishing pyTFM, which was crucial to accomplishing the study this protocol refers to.
Competing interests
There is no conflict of interest we are aware of.
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Article Information
Publication history
Received: Jun 3, 2024
Accepted: Oct 29, 2024
Available online: Nov 21, 2024
Published: Jan 5, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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5,157 | https://bio-protocol.org/en/bpdetail?id=5157&type=0 | # Bio-Protocol Content
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Protocol for Immune Cell Isolation, Organoid Generation, and Co-culture Establishment from Cryopreserved Whole Human Intestine
EG Enrique Gamero-Estevez *
IH Inga Viktoria Hensel *
MS Michelle Steinhauer *
OM Olivia Müllertz
ES Elizaveta Savochkina
IS Ibrahim Murathan Sektioglu
BS Bilgenaz Stoll
SD Shaghayegh Derakhshani
SD Sarah Devriese
KK Kyungbo Kim
MR Martin Resnik-Docampo
(*contributed equally to this work)
Published: Vol 15, Iss 1, Jan 5, 2025
DOI: 10.21769/BioProtoc.5157 Views: 730
Reviewed by: Andrea GramaticaPhilipp Wörsdörfer Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in EMBO Molecular Medicine Mar 2024
Abstract
The human intestine plays a pivotal role in nutrient absorption and immune system regulation. Along the longitudinal axis, cell-type composition changes to meet the varying functional requirements. Therefore, our protocol focuses on the processing of the whole human intestine to facilitate the analysis of region-specific characteristics such as tissue architecture and changes in cell populations. We describe how to generate a biobank that can be used to isolate specific immune cell subtypes, generate organoid lines, and establish autologous immune cell-organoid co-cultures.
Key features
• Dissection and tissue analysis of whole human intestines.
• Cryopreservation for biobank generation.
• Optimized protocols for the isolation of epithelial and immune cells.
• Autologous co-culture of organoids and lamina propria–derived immune cells.
Keywords: Whole gut collection Tissue processing Biobank Cryopreservation Immune cell isolation Intestinal organoid Autologous co-culture Lamina propria Intestinal immune organoids
Graphical overview
Human intestine processing to cryopreserve biopsies, isolate epithelial and immune cells, and generate organoids and autologous co-culture. Created with BioRender.com.
Background
The intestine is the longest organ of the human body and possesses the largest surface area, being also constantly exposed to commensal and pathogenic bacteria. Thus, maintaining barrier integrity and accurate immune surveillance is crucial to preserve its function and homeostasis. A dysfunction in the epithelial barrier or in the immune cell composition of the intestine can lead to severe and chronic diseases, from cancer to autoimmune diseases. Therefore, it is essential to understand the interaction between the epithelial barrier and the immune cells as well as to identify region-specific immune cell subpopulations.
Access to intestinal tissue is limited, and most research relies on small biopsies (<5 cm) or intestinal resections (<40 cm long) [1–3]. However, to fully understand the complexity and regional variations, studying the intestine in its entirety is necessary. Additionally, this allows the analysis of the jejunum and ileum, which are particularly difficult to address with biopsies or resections.
Processing such a large organ requires a stringent protocol that focuses on the cryopreservation and time-independent analysis of the samples. Moreover, the focus of most cryopreservation methods is on the recovery of a specific cell population [4,5], but they do not address the isolation of cells from different intestinal regions or simultaneous epithelial and immune cell isolation [3,6].
Our protocol closes this gap by describing a cryopreservation method that supports the preservation of both immune and epithelial cells of various intestinal regions while additionally being applicable to cell isolations from fresh tissue. It serves as an essential and versatile guideline for the preservation and cell isolation of the whole human intestine while providing additional options for tissue analysis.
We give a detailed description of an optimized and refined protocol that describes how to process the whole intestine, generate a biobank, and perform cell isolation from fresh and cryopreserved tissue. In particular, techniques for the generation of intestinal organoids and the isolation of epithelium and immune cells with enrichment of specific cell fractions depending on the enzyme digestion carried out are presented. Finally, we provide a protocol to establish an autologous immune cell-organoid co-culture, which can be further modified to study one specific intestinal region or immune cell subtype.
Materials and reagents
Biological materials
1. Human whole intestinal tissues (Novobiosis, I3PT biobank, DTI Foundation)
2. R-Spondin 1 cell line, stably transfected HEK293T cells (kindly provided by Hugo de Jonge)
Reagents
General buffers and media
1. Sodium chloride (NaCl) (Carl Roth, catalog number: 3957.2)
2. Potassium di-hydrogen phosphate (KH2PO4) (Carl Roth, catalog number: P018.2)
3. Di-sodium hydrogen phosphate dihydrate (Na2HPO4·2H2O) (Carl Roth, catalog number: T877.1)
4. Heat-inactivated FBS (Capricorn Scientific, catalog number: FBS-HI-11A)
5. PBS-/- (Capricorn Scientific, catalog number: PBS-1A)
6. HEPES (Capricorn Scientific, catalog number: HEP-B)
7. 0.5 M EDTA pH 8.0 (Fisher Scientific, catalog number: 46-034-CI)
8. Sucrose for molecular biology, ≥99.5% (GC) (Sigma-Aldrich, catalog number: S0389-500G)
9. RPMI 1640 medium, 500 mL (Thermo Fisher Scientific, catalog number: 21875034)
10. Custodiol (Dr. Franz Köhler Chemie)
11. DTT (1,4-Dithiothreitol) (Carl Roth, catalog number: 6908.3)
12. Ethanol Rotipuran ≥99.8% p.a. (Carl Roth, catalog number: 9065.05)
13. Methanol (Carl Roth, catalog number: 8388.6)
Buffers and media for cryopreservation
1. Isopropanol 99.8% (neoLab Migge, catalog number: 2-3701)
2. CryoSure-DMSO USP grade (WAK-Chemie Medical, catalog number: WAK-DMSO-10)
3. Ethylene glycol anhydrous, 99.8% (Sigma-Aldrich, catalog number: 324558-100ml)
4. D-(+)-trehalose dihydrate (Sigma-Aldrich, catalog number: T0167-25G)
5. RNA later stabilization solution (Qiagen, Fisher Scientific, catalog number: AM7021)
6. CryoStor® CS10 (STEMCELL Technologies, catalog number: 07930)
Chemicals for histology
1. FSC 22 frozen section media, blue, 9 × 118 mL (Leica, catalog number: 3801481)
2. Paraformaldehyde solution 4% in PBS (Santa Cruz Biotechnology, catalog number: sc-281692)
3. Paraplast (Paraffin) (Carl Roth, catalog number: X880.2)
4. Xylene (Sigma-Aldrich, catalog number: 534056)
5. Tissue freezing medium (Leica, catalog number: 14020108926)
Buffers and media for organoid generation
1. Matrigel growth factor reduced (GFR) basement membrane matrix, phenol red-free, LDEV-free (Corning, Fisher Scientific, catalog number: 356231)
2. StemProTM AccutaseTM cell dissociation reagent (Thermo Fisher, Gibco, catalog number: A1110501)
3. Advanced DMEM/F12 medium (Fisher Scientific, Gibco, catalog number: 12634010)
4. GlutaMAX (Thermo Fisher Scientific, catalog number: 35050038)
5. Primocin (Invivogen, catalog number: ant-pm-1)
6. B-27 supplement (Thermo Fisher Scientific, catalog number: 17504044)
7. WNT surrogate-Fc fusion protein (ImmunoPrecise, catalog number: N001)
8. R-Spondin 1-conditioned medium (produced by R-Spondin 1 cell line)
9. n-Acetyl-cysteine (Sigma-Aldrich, catalog number: A7250)
10. Nicotinamide (Sigma-Aldrich, catalog number: N0636-500G)
11. Recombinant murine EGF (PeproTech, catalog number: 315-09)
12. Murine noggin (Peprotech, catalog number: 250-38)
13. Recombinant human IGF-1 (PeproTech, catalog number: 100-11)
14. Recombinant human FGF-2 (PeproTech, catalog number: 100-18C)
15. Human Leu[15]-Gastrin I (Sigma-Aldrich, catalog number: G9145)
16. A83-01 (Sigma-Aldrich, catalog number: SML0788)
17. SB202190 (Tocris, catalog number: 1264)
18. Y-27632 (Hölzel Diagnostika, catalog number: HY-10583)
Buffers and media for cell isolation
1. HBSS without Ca2+/Mg2+ (Thermo Fisher Scientific, catalog number: 14175129)
2. Multi Tissue Dissociation kit (Miltenyi BioTec, catalog number: 130-110-201)
3. Collagenase type IV (Fisher Scientific, catalog number: 17104019)
4. DNase I (Sigma-Aldrich, Chemie, catalog number: 10104159001)
5. LiberaseTM TL research grade (Sigma-Aldrich, catalog number: 5401020001)
Flow cytometric analysis
1. CD45 BV510 (BD Biosciences, catalog number: 563204)
2. EPCAM (CD326) PE (BioLegend, catalog number: 324206)
3. CD3 BUV496 (BD Biosciences, catalog number: 612940)
4. CD19 BV750 (BioLegend, catalog number: 302262)
5. HLADR BUV395 (BD Biosciences, catalog number: 564040)
6. CD11c BB515 (BD Biosciences, catalog number: 564490)
7. CD14 BUV737 (BD Biosciences, catalog number: 612763)
8. CD56 APC/Cy7 (BioLegend, catalog number: 318332)
9. CD103 APC (BioLegend, catalog number: 350216)
10. 7AAD (BioLegend, catalog number: 420404)
Supplies for co-culture
1. Percoll (Thermo Fisher Scientific, catalog number: 17-0891-01)
2. Glutaraldehyde 50% (Sigma-Aldrich, Chemie, catalog number: G7776-10ML)
3. Formaldehyde solution 37% (Carl Roth, catalog number: CP10.1)
Solutions
1. 10× phosphate-buffered saline (PBS) (see Recipes)
2. Cryopreservation buffer (see Recipes)
3. Complete Roswell Park Memorial Institute medium (cRPMI) (see Recipes)
4. Mucus removal solution (see Recipes)
5. 30% ethanol (see Recipes)
6. 50% ethanol (see Recipes)
7. 70% ethanol (see Recipes)
8. 80% ethanol (see Recipes)
9. 5% sucrose (see Recipes)
10. 15% sucrose (see Recipes)
11. 30% sucrose (see Recipes)
12. Dissociation solution (see Recipes)
13. Base medium (see Recipes)
14. Growth medium (see Recipes)
15. Expansion medium (see Recipes)
16. 0.75 M sucrose (see Recipes)
17. 0.375 M sucrose (see Recipes)
18. 0.125 M sucrose (see Recipes)
19. 10% FBS/PBS (see Recipes)
20. Epithelial cell removal media (see Recipes)
21. Enzymatic mixture A (see Recipes)
22. Enzymatic mixture B (see Recipes)
23. Enzymatic mixture C (see Recipes)
24. 80% Percoll (see Recipes)
25. 40% Percoll (see Recipes)
26. Co-culture fixative (see Recipes)
Recipes
All buffers can be stored at room temperature unless otherwise specified. All sucrose buffers can be stored at 4 °C and used within 4 weeks.
1. 10× phosphate-buffered saline (PBS)
Reagent Final concentration Amount
NaCl 1.37 M 80.063 g
KH2PO4 0.018 M 2.45 g
Na2HPO4·2H2O 0.103 M 18.333 g
H2O n/a 900 mL
Total 1 L
Adjust the pH of 1× PBS to 7.4.
2. Cryopreservation buffer
Adapted from Hughes et al. [4]. Store at 4 °C and use within 72 h.
Reagent Final concentration Amount
D-(+)-Trehalose dihydrate 50 mM 3.78 g
CryoSure-DMSO USP grade 6% 12 mL
Ethylene glycol anhydrous, 99.8% 5% 10 mL
Heat-inactivated FBS 178 mL
Total 200 mL
3. Complete Roswell Park Memorial Institute medium (cRPMI)
Store at 4 °C and use within 4 weeks.
Reagent Final concentration Amount
RPMI n/a 450 mL
FBS 5% 50 mL
Total 500 mL
4. Mucus removal solution
Store at 4 °C and use within 12 h.
Reagent Final concentration Amount
RPMI n/a 450 mL
FBS 5% 50 mL
DTT (100%) 4 mM 0.3 g
Total 500 mL
5. 30% ethanol
Reagent Final concentration Amount
Ethanol (absolute) 30% 300 mL
H2O n/a 700 mL
Total 1,000 mL
6. 50% ethanol
Reagent Final concentration Amount
Ethanol (absolute) 50% 500 mL
H2O n/a 500 mL
Total 1,000 mL
7. 70% ethanol
Reagent Final concentration Amount
Ethanol (absolute) 70% 700 mL
H2O n/a 300 mL
Total 1,000 mL
8. 80% ethanol
Reagent Final concentration Amount
Ethanol (absolute) 80% 800 mL
H2O n/a 200 mL
Total 1,000 mL
9. 5% sucrose
Reagent Final concentration Amount
Sucrose 5% 5 g
PBS n/a 100 mL
Total 100 mL
10. 15% sucrose
Reagent Final concentration Amount
Sucrose 15% 15 g
PBS n/a 100 mL
Total 100 mL
11. 30% sucrose
Reagent Final concentration Amount
Sucrose 30% 30 g
PBS n/a 100 mL
Total 100 mL
12. Dissociation solution
Reagent Final concentration Amount
PBS-/- n/a 9.8 mL
EDTA (0.5 M) 10 mM 200 μL
Total 10 mL
13. Base medium
Store at 4 °C and use within 4 weeks.
Reagent Final concentration Amount
Advanced DMEM/F12 n/a 489 mL
HEPES 10 mM 5 mL
GlutaMAX (100×) 1× 5 mL
Primocin 100 μg/mL 1 mL
Total 500 mL
14. Growth medium
Store at 4 °C and use within 2 weeks.
Reagent Final concentration Amount
Advanced DMEM/F12 n/a 6.5379 mL
R-Spondin 1-conditioned medium 20% 2 mL
B-27 supplement 1× 200 μL
HEPES 10 mM 100 μL
GlutaMAX (100×) 1× 100 μL
Nicotinamide 10 mM 100 μL
Murine noggin 100 ng/mL 50 μL
N-acetyl-cysteine 1.25 mM 25 μL
Primocin 100 μg/mL 20 μL
Surrogate WNT 0.5 nM 10 μL
EGF 50 ng/mL 10 μL
SB202190 10 μM 3.3 μL
A83-01 500 nM 1 μL
Human Leu[15]-Gastrin I 10 nM 1 μL
Y-27632* 10 μM 1 μL
Total 10 mL
*First 2–3 days after isolation or expansion.
15. Expansion medium
Store at 4 °C and use within 2 weeks.
Reagent Final concentration Amount
Advanced DMEM/F12 n/a 7.472 mL
R-Spondin 1-conditioned medium 20% 2 mL
B-27 supplement 1× 200 μL
HEPES 10 mM 100 μL
GlutaMAX (100×) 1× 100 μL
Murine noggin 100 ng/mL 50 μL
N-acetyl-cysteine 1 mM 20 μL
Primocin 100 μg/mL 20 μL
Surrogate WNT 0.5 nM 10 μL
EGF 50 ng/mL 10 μL
Recombinant human IGF-1 100 ng/mL 10 μL
Recombinant human FGF-2 50 ng/mL 5 μL
A83-01 500 nM 1 μL
Human Leu[15]-Gastrin I 10 nM 1 μL
Y-27632* 10 μM 1 μL
Total 10 mL
*First 2–3 days after isolation or expansion.
16. 0.75 M sucrose
Reagent Final concentration Amount
Sucrose 0.75 M 5.13 g
PBS 90% 18 mL
FBS 10% 2 mL
Total 20 mL
17. 0.375 M sucrose
Reagent Final concentration Amount
Sucrose 0.375 M 2.6 g
PBS 90% 18 mL
FBS 10% 2 mL
Total 20 mL
18. 0.125 M sucrose
Reagent Final concentration Amount
Sucrose 0.125 M 0.9 g
PBS 90% 18 mL
FBS 10% 2 mL
Total 20mL
19. 10% FBS/PBS
Store at 4 °C and use within 72 h.
Reagent Final concentration Amount
PBS 90% 36 mL
FBS 10% 4 mL
Total 40 mL
20. Epithelial cell removal media
Store at 4 °C and use within 72 h.
Reagent Final concentration Amount
HBSS (-/-) n/a 39.5 mL
HEPES (1 M) 100 mM 5 mL
FBS 10% 5 mL
EDTA (0.5 M) 5 mM 500 μL
Total 50 mL
21. Enzymatic mixture A
Prepare directly before use.
Reagent Final concentration Amount
cRPMI n/a 4.725 mL
Liberase (2.5 mg/mL) 100 μg/mL 200 μL
DNase (10 mg/mL) 150 μg/mL 75 μL
Total 5 mL
22. Enzymatic mixture B
Prepare directly before use.
Reagent Final concentration Amount
cRPMI n/a 4.89 mL
Collagenase IV (0.327 g/mL) 2 mg/mL 31 μL
DNase (10 mg/mL) 150 μg/mL 75 μL
Total 5 mL
23. Enzymatic mixture C (Multi Tissue Dissociation kit 1: MTKD1)
Prepare directly before use.
Reagent Final concentration Amount
cRPMI n/a 4.675 mL
Enzyme A (MTDK1) n/a 25 μL
Enzyme R (MTDK1) n/a 100 μL
Enzyme D (MTDK1) n/a 200 μL
Total 5 mL
24. 80% Percoll
Store at 4 °C and use within 4 weeks.
Reagent Final concentration Amount
Percoll (100%) 80% 72 mL
10× PBS n/a 8 mL
PBS-/- n/a 20 mL
Total 100 mL
25. 40% Percoll
Store at 4 °C and use within 4 weeks.
Reagent Final concentration Amount
Percoll (100%) 80% 54 mL
10× PBS n/a 6 mL
PBS-/- n/a 40 mL
Total 100 mL
26. Co-culture fixative
Prepare on the day of use.
Reagent Final concentration Amount
Paraformaldehyde solution 37% in PBS 3.7% 1 mL
Glutaraldehyde 50% 0.1% 20 μL
PBS-/- 8.98 mL
Total 10 mL
Laboratory supplies
General
1. 15 mL tubes (neoLab Migge, catalog number: 352096)
2. 50 mL tubes (neoLab Migge, catalog number: 352070)
3. 5 mL serological pipette (Falcon, neoLab Migge, catalog number: 357543)
4. 10 mL serological pipette (Falcon, neoLab Migge, catalog number: 357551)
5. 25 mL serological pipette (Falcon, neoLab Migge, catalog number: 357525)
6. Cell strainer 100 μm (Greiner Bio-One, catalog number: 542000)
7. Cell strainer 70 μm (Greiner Bio-One, catalog number: 542070)
8. FACS tubes, Falcon 5 mL round-bottom polystyrene test tube, with cell strainer snap cap (Fisher Scientific, catalog number: 352235)
9. SafeSeal tubes 1.5 mL (Sarstedt, catalog number: 72.706)
Supplies for lab preparation
1. Autoclaving bags, large size (Greiner Bio-One, catalog number: 649201)
2. Labeling tape (neoLab Migge, catalog number: 2-6101)
3. Sterile drape Absorba-II, 200 × 320 cm (Integra, catalog number: 2206-3114)
4. Plastic tray (neoLab Migge, catalog number: 2-3853)
5. Forceps, small (Vubu Medical Instruments, catalog number: VUBU-02-45911 and VUBU-02-8911)
6. Forceps, big (Vubu Medical Instruments, catalog number: VUBU-02-21921 and VUBU-02-67920)
7. Intestine scissors (Vubu Medical Instruments, catalog number: VUBU-03-30521)
8. Scissors, curved (Vubu Medical Instruments, catalog number: VUBU-03-16114)
9. Scissor small (Vubu Medical Instruments, catalog number: VUBU-03-44908)
10. Petri dish 9 cm (Carl Roth, catalog number: N221.2)
11. Cooling element, Va-Q-accu 27L +05G (Va-Q-tec, catalog number: AK000295)
12. Ice bucket with lid (neoLab, catalog number: 2-6002)
13. Dry ice nuggets (CLEANGAS)
14. Measuring tape (Amazon, BLAMT, Stanley, catalog number: 0-34-295)
15. Safety pin 4.5 cm (Amazon, TK Gruppe Timo Kringler, catalog number: B082ZQZTDQ)
16. Wide metal dish 29 × 23 × 4 cm (Rewe, catalog number: 32420)
17. Bench cover, Foliodrape protect cover cloth 45 × 75 cm (Medpex, catalog number: 798179)
18. Labcoat (neoLab Migge, catalog number: 4-1482)
19. Protective gowns (Carl Roth, catalog number: CAL9.1)
20. Sleeves Medi-Inn (Carl Roth, catalog number: 1PP4.1)
21. Plastic box 15.7 × 11.8 × 10.0 cm, 1 L (Amazon, Rotho, catalog number: 6116705070)
Supplies for tissue collection
1. Freezing container, 18 × 1–2 mL (neoLab Migge, catalog number: 2-3701)
2. Barcoded cryovials, 2 mL (Greiner Bio-One, catalog number: 122280-2DG)
3. Scanner (CSL-Computer, catalog number: 722304674722 via Amazon)
4. Black cryo rack (Greiner Bio-One, catalog number: 802576)
5. White cryo rack (Greiner Bio-One, catalog number: 802202)
6. Beaker 800 mL (neoLab Migge, catalog number: E1052)
7. Sealing film parafilm M, 100 mm, 38 m (Carl Roth, catalog number: H666.1)
8. S-Monovette Serum-Gel 7.5 mL (Sarstedt AG & Co. KG, catalog number: 01.1602)
9. Cotton swaps, sterile (P+W Medizintechnik Platz & Co Handels GmbH, catalog number: 10724)
Supplies for histology
1. Histology cassettes (neoLab Migge, catalog number: 70012)
2. TT Cryomold intermediate, 15 × 15 × 15 mm (Science Service, catalog number: SA62534-15)
3. BD Microlance 3 Nr. 20, 27 G needle (MSG Medizinische Geräte, Handel und Service, catalog number: BD302200)
Supplies for organoid generation
1. 24-well plate (Greiner Bio-One, catalog number: 662160)
2. 200 μL pipette tips without filter (Sarstedt, catalog number: 70.760.202)
Supplies for immune cell isolation
1. Cell strainer 40 μm (Greiner Bio-One, neoLab Migge, catalog number: 352340)
2. 35 mL GosselinTM X500 FIOLE DROSO 25X95MM VRAC PS PS (Fisher Scientific, catalog number: 11773303)
3. Magnetic stirring bar, PTFE, cylindrical (BRAND, Merck Chemicals, catalog number: Z328723-10EA)
4. Neubauer counting chamber (neoLab Migge, catalog number: 191844172)
5. Duran crystallizing beaker (Merck Chemicals GmbH, catalog number: Z231754)
6. C-tubes for gentleMACS (Miltenyi BioTec, catalog number: 130-093-237)
7. Round-bottom 96-well plates (neoLab Migge, catalog number: C-8207)
Equipment
1. Pipet boy acu 2 (Integra BioscienceTM, catalog number: 10060262)
2. Cell culture incubator (Heracell, catalog number: 51032720)
3. Herasafe 2030i biosafety hood (Thermo Fisher Scientific, catalog number: 51032324)
4. Forced convection laboratory incubator (ScanCell, catalog number: 37)
5. 5810 R centrifuge (Eppendorf, catalog number: 5811000420)
6. Fluorescence microscope (Nikon, model: Eclipse Ti2)
7. Microscope stage incubator (Tokai Hit)
8. Rocker shaker 2D series (Carl Roth GmbH+Co. KG, catalog number: XT47.1)
9. IKA 4011000 Roller shaker (IKA, catalog number: EW-04304-05)
10. CryoStar NX50 (Thermo Fisher Scientific, EprediaTM, catalog number: 13435949)
11. HM 340E Microtome (Thermo Fischer Scientific, catalog number: 26370735)
12. gentleMACS Octo Dissociator with Heaters (Miltenyi Biotec B.V., catalog number: 130-096-427)
13. Spectral Analyzer (SONY, model: ID7000TM)
14. Electrothermal Paraffin section flotation bath (Cole-Parmer, catalog number: MH8517)
15. Orbital shaker (Sunlab, catalog number: SU1030)
16. Scale (Sartorious, catalog number: CP32025-OCE)
17. Vortexer (Neolab, catalog number: D6012)
18. -80 °C freezer
19. -20 °C freezer
20. 4 °C fridge
21. N2 tank
22. Paraffin processing station: Leica, ASP 300S Tissue Processor
23. Casting station: Leica, HistoCore Arcadia H Embedding Center
24. Magnetic stirrer with heater (IKA, model: RCT5 digital)
Software and datasets
1. FlowJo v10.8.1
2. GraphPad Prism v.10.0.0
3. Fiji (ImageJ)
4. Microsoft Excel
Procedure
The protocol below describes the processing and generation of a tissue biobank as well as the isolation of immune and epithelial cells from both fresh and cryopreserved tissue. More specifically, it describes the downstream analysis of the tissue itself, including histology, RNA, and protein isolation. It addresses the in-depth analysis of immune cell subtypes using flow cytometry and cell sorting. Additionally, it provides a guide for establishing region-specific organoids, which can be combined with isolated immune cells to generate an autologous co-culture. Possible readouts of the co-culture include cell interaction studies, cytokine release assays, and single-cell RNA sequencing. This extensive protocol can be customized to individual requirements, e.g., the number and location of collected biopsies, as well as the choice of specific downstream applications.
A. Protocol for surgeons
This section describes the established protocol for organ procurement.
1. Whole organ processing prior to shipment
a. After excising the whole intestine distal to the pylorus and at the rectum, separate the small and large intestine at the ileocecal valve.
b. Label the proximal and distal sides.
Note: Several options can be applied to improve tissue integrity. A small incision can be made along the longitudinal axis, the mesentery removed, and the intestine cut open. Additionally, Custodiol can be used to flush and remove luminal content.
c. Transfer the organ to Custodiol and package and store it on ice similar to any other donor organ.
d. Draw blood and store it in a serum or plasma tube, depending on the desired readout.
e. Send it to the recipient.
B. Region definition
This section describes how the intestinal regions are defined.
Extensive literature research was performed to define the different small and large intestinal regions. Since the absolute length of the intestine varies between individuals, we decided to standardize the sampling location based on relative values. We performed a meta-analysis to define average values for total length and to identify the different gut regions [7–12]. Based on these values, we calculated relative values for the beginning and end of the six regions: duodenum, jejunum, and ileum, as well as the ascending, transverse, and descending colon, defining the proximal and the distal end as 100% for either the small or large intestine, respectively. Figure 1 shows the six chosen regions and their definition with the ileocecal valve as a reference. Small intestinal regions are marked with negative relative values and large intestinal regions with positive values. The absolute values can then be calculated for the individual intestine using the total length of the small and large intestine, respectively (Supplemental Excel file).
Figure 1. Regional description of the human gut. Small intestine (pink) and colon (brown) are separated by regions depending on their relative distance to the ileocecal valve (reference point 0%) as a reference point. Approximate lengths of the small and large intestine are shown [10,12]. Percentages use the total length of either the small or large intestine as a reference. A negative percentage indicates the distal to proximal orientation in the case of the small intestine.
C. Lab preparation
This section gives details about required lab preparations, the establishment of a biobank file, and workflow. The following steps should be carried out a day before receiving the gut.
1. Biobank preparation
a. Generate a biobank file using the template (Supplemental Excel file) containing information on the date, patient age, sex, and medical history if provided. Adjust the file according to the regions sampled, number of tubes collected, and types of storage (biobank, RNA, protein, or histology).
b. Label white cryo racks according to the gut ID.
c. Label the freezing containers according to each gut region and, if necessary, number them inside to track the tube order easily (Figure 2A). Fill it with isopropanol to the indicated level.
Figure 2. Laboratory preparation a day before the arrival of the organ. (A) Freezing containers labeled per gut region and filled with barcoded cryotubes. (B) An example of bench coverage using autoclavable bags and drapes.
d. Transfer the barcoded cryovials to the freezing container.
Note: Regular cryovials can be used if no biobank is generated or tubes are labeled instead of the barcode.
e. Prepare the cryopreservation buffer. The final volume required can be calculated by the total number of tubes × 1 mL/tube.
f. Pipette 1 mL of the cryopreservation buffer to each 2 mL cryovial using a 10 mL pipette.
g. Enter the barcode of each tube in the biobank file according to its location in the freezing container. This can be done using a barcode scanner.
h. Transfer the freezing containers to 4 °C (Figure 2A).
2. Tubes for mucosal sampling, blood, RNA, and protein isolation
a. For mucosal sampling, prepare empty cryovials by labeling them with M1–M6.
b. For blood (serum/plasma) storage, prepare the desired number of empty tubes labeled with B1–X.
c. For RNA isolation, fill a barcoded cryovial with 1 mL of RNA later stabilization solution.
d. Prepare empty barcoded cryovials for protein isolation.
e. Transfer all of them to a black cryo rack labeled accordingly (e.g., Box 3).
Note: Since the tubes will be taken out of the box for sample transferal, it is advisable to label the lid with the region number and type of storage (RNA or protein) to ease the sampling process.
f. Enter the barcode of each tube in the biobank file according to its location in the cryo rack.
Note: Print a hard copy of the biobank file to make it easy to track the barcode back to the intestinal region during tissue processing.
g. Label two beakers with paraffin or sucrose, respectively, and fill them with 300 mL of 4% PFA. Seal with Parafilm and store at 4 °C.
h. Label the histology cassettes and cryomolds.
i. If a larger tissue should be fixed as a Swiss roll, label 27 G needles accordingly.
3. Laboratory preparation
Note: It is advisable to remove any equipment and material from the workbenches beforehand.
a. Cover benches with plastic cover (e.g., cut-open large autoclaving bags) and fix them on the benches (Figure 2B).
b. Use the sterile drape to add another protective layer on the benches (Figure 2B).
c. Prepare one tray with all the equipment required to dissect the tissue, including scissors, tweezers, two 27 G needles, one rack for cryovials, and one Petri dish. The workspace can be optionally equipped with a lamp.
d. Prepare a tray with measuring tape, dissecting scissors, and safety pins necessary for initial tissue measurement and labeling.
d. Print a workflow for each region containing all the steps that need to be followed.
Note: This will help you go through the sampling process and allow you to take notes on tissue integrity.
e. Transfer the cooling elements to -20 °C.
f. Prepare two bins with autoclaving bags—one for general waste and one designated only for intestinal tissue waste.
Note: Any residual tissue needs to be discarded following the rules for human tissue disposal.
D. Whole gut tissue processing
This section describes the processing of the procured gut tissue and how it is divided into smaller pieces for the respective intestinal regions.
1. Gut receival.
One hour before
a. Prepare one ice box per region to be sampled.
b. In case isolation from fresh tissue for either immune or epithelial cells is desired, prepare a 15 mL tube containing Custodiol or RPMI and place it on ice.
c. Transfer the barcoded cryovials for RNA and protein samples prepared the day before to the ice box for each region, respectively.
d. Prepare a box with dry ice and place the empty cryo rack inside for RNA and protein sampling.
e. Prewarm the mucus removal solution and the cRPMI at 37 °C.
Upon delivery
Critical: You are handling human material. Therefore, it is important to wear personal safety equipment. The staff handling the tissue are advised to use extra impermeable sleeves.
Note: The tasks should be distributed beforehand amongst the team members and should be carried out side-by-side.
f. Prepare an empty big plastic tray located next to the sink.
Note: Depending on the delivery conditions, the ice contained in the box can be completely melted. Be aware that there can be water spillage.
g. Open the carton, cut open the first plastic bag, and retrieve the Styrofoam box. The intestine will be packed in two additional plastic bags or containers.
Note: The tissue will often come separated by small intestine and colon. If not, the first step would be to separate them by cutting through the ileocecal valve (Figure 1). Then, you should start with the region of interest while the other can stand in Custodiol until it is ready for processing (usually after step H10).
h. Place the plastic bag containing the intestine in the plastic tray and cut it open.
i. Remove any residual liquid.
j. Rinse the tissue with cold PBS if intestinal content has spilled. Large quantities of stool can be removed manually.
Note: Steps D1j depend on the individual preprocessing of the gut. If little or no intestinal content is present or if the intestine is cut open, move to the next step.
2. Measurement, region identification, and tissue section preparation
a. Place the tissue on the prepared bench and identify the proximal and distal ends (Figure 3A).
Note: Depending on the preprocessing, missing part of the intestine or a lack of labeling of the openings can be challenging. Familiarize yourself beforehand with the anatomical features of the small and large intestines. The duodenum is free of mesentery, and the folding of the mucosa is more pronounced compared to the other intestinal segments. Additionally, the entry of the bile duct might be identifiable. The color of the mucosal lining is more pinkish red in the ileum than in the duodenum when directly compared to each other. For the large intestine, the appendix and cecum can be used as hallmarks to identify the proximal end. Additionally, depending on the excision site, the rectum can be identified at the distal end by longitudinal striation.
Figure 3. Gut processing and dissection. (A) Comparison of the proximal and distal ends of the small intestinal segments: ileum (left) and duodenum (right). (B) Removing the mesentery, cutting it as close to the tissue as possible to allow proper measurements. (C) Spreading the tissue on the table to avoid wrinkles or stretches to measure its length. (D) Measurement of the large intestine. (E) Example of the excision of a 40 cm piece of ileum for biopsy collection and a 1 cm piece for histology. (F) Longitudinal opening of the 40 cm ileal piece for further dissection.
b. Once the proximal and distal ends are identified, the mesentery is removed (Figure 3B).
Note: Excise as close to the mucosal tissue as possible. This will facilitate tissue spreading to its full length.
c. Spread out the tissue to its full length without stretching it (Figure 3C).
d. Measure the total length of the tissue using a measuring tape and document it in the biobank file (Figure 3D).
Note: In the template, the incision sites will be calculated in centimeters once the total length is added.
e. Make an incision at the calculated length and make another incision 20–30 cm toward the proximal (small intestine) or distal (large intestine) opening (Figure 3E).
f. Mark the distal end with a safety pin.
g. Cut one 1 cm piece of the proximal side for histology and follow step G1 (Figure 3E).
h. Open the intestinal piece longitudinally by cutting following the mesentery (Figure 3F).
i. Continue on section H.
E. Blood preparation for storage
This section below is optional and depends on the individual biobanking desires. The blood is collected in serum or plasma tubes from the patient by the surgeon. The collected blood needs to be processed according to the desired downstream experiments. Potentially blood-derived immune cells could be isolated, analyzed, or used as an alternative source for co-culture. The protocol below describes the collection of serum from blood collected in serum tubes that promote blood coagulation.
1. Place the tube containing the blood into the pre-cooled centrifuge and spin down at 750× g for 15 min.
2. Transfer the supernatant-containing serum to 1.5 mL tubes labeled B1–X.
3. Transfer the tubes to the designated box on dry ice and store them at -80 °C.
Note: After organoid establishment, it can then be used in an organoid-serum stimulation assay, as shown in a novel Systemic Lupus Erythematosus (SLE) model [13].
F. Mucus collection for microbiome analysis
This is an optional step that can be performed if microbiome analysis is desired.
1. Make sure the mucosal side of the tissue is facing upward.
2. Use a sterile cotton swap and sample the mucus by wiping and rolling it over the tissue piece.
3. Transfer the swap to an empty 1.5 mL tube labeled with M1–M6 and break off the handle.
4. Transfer the tube to the designated box on dry ice and store at -80 °C.
G. Tissue collection for histology
This section gives a detailed description of tissue processing if histological staining is desired.
1. Tissue fixation
a. Cut the tissue from step D2g into pieces of the desired size and rinse with Custodiol. If a Swiss roll is desired, open the tissue longitudinally, roll the tissue, and fix it with a needle.
b. Transfer the tissues into the tissue cassettes and drop them into the respective prepared beaker containing 4% PFA (prepared on step C2g).
Critical: Make sure that the tissue is placed flat into the cassette and that the lid does not squeeze the tissue to ensure undisturbed tissue integrity.
Note: Best villi preservation is obtained if the intestinal biopsy is not open and remains closed.
Note: Swiss rolls are a good alternative if tissue has been cut open. Consider fixing it in a 50 mL tube and then transferring it into the cassette after fixation to improve villi and tissue histology.
c. Fix the tissue overnight at 4 °C.
2. Embedding of fixed tissue for paraffin sections
Note: All following steps are performed at room temperature except stated otherwise.
a. Transfer the cassettes with tissue to 30% ethanol for 1 h.
b. Transfer the cassettes to 50% ethanol for 1 h.
c. Transfer the cassettes to 70% ethanol for 1 h.
Note: Cassettes can be kept in 70% ethanol at 4 °C for up to a month before continuing to the next steps.
d. Transfer the cassettes to 80% ethanol for 1 h.
e. Transfer the cassettes to 90% ethanol for 30 min.
f. Transfer the cassettes to 100% ethanol for 90 min.
Note: If slower dehydration is desired, more intermediate steps can be added using 95% and 98% ethanol.
g. Transfer the cassettes to xylene for 1 h.
h. Transfer the cassettes to fresh xylene for 1 h.
i. Transfer the cassettes to melted paraffin for 1 h at 65 °C.
j. Transfer the cassettes to fresh melted paraffin for at least 3 h at 65 °C.
Note: Steps G2a–j can be performed in an automated paraffin processing station.
k. Remove the tissue from the cassettes and embed them using a casting station to ensure the desired histology orientation.
l. Let the cassettes cool down and store them at either room temperature or at 4 °C.
3. Embedding of fixed tissue for cryosections
Note: All following steps are performed at room temperature except stated otherwise.
a. Transfer the cassettes to 5% sucrose for 1 h.
b. Transfer the cassettes to 15% sucrose for 1 h.
c. Transfer the cassettes to 30% sucrose for 1 h.
Note: It is recommended to leave the tissue in 30% sucrose overnight at 4 °C. The tissue can be stored in 30% sucrose indefinitely, but it is recommended to be processed fast since contamination may appear.
d. Transfer the cassettes to a mix of 50:50 of tissue freezing medium and 30% sucrose for 1 h.
e. Place the tissues in cryomolds filled with tissue freezing medium and position the tissue.
f. Freeze the cryomolds containing the tissue by dropping them on liquid nitrogen or in a methanol-dry ice bath.
g. Wrap the cryomolds in aluminum foil and store at -80 °C.
H. Mucus removal
This section describes how to remove the secreted mucus from the underlying tissue. This step is essential to improve cell recovery yields.
Note: Continue from step D2i.
1. Transfer the tissue piece to a plastic box labeled with the designated intestinal area and containing 150 mL of Custodiol.
2. Transfer it to an ice box.
Note: The transfer to Custodiol and ice is necessary to both add a cleaning step to remove debris and to ensure the tissue is preserved during further processing.
3. Add 100 mL of prewarmed mucus removal solution into a wide metal or plastic flat-bottom dish at room temperature.
Note: Depending on the container size, a larger volume of solution may be needed. Ensure that the tissue is fully covered.
4. Transfer the open and clean piece of tissue into the wide metal or plastic flat bottom dish with the mucosa facing toward the bottom of the dish.
Note: It is recommended to use pieces of 20 cm to avoid excessive or uneven treatment. If larger pieces are desired, it is better to do it sequentially, 20 cm at a time.
5. Gently press and shake the tissue with forceps against the bottom of the dish for 10 min.
Note: This step is important since the gentle friction against the bottom of the container will help the mucus to detach.
Critical: Too much pressure will lead to tissue damage, while too little will not remove big pieces of mucus effectively.
6. Remove the mucus removal solution and add 100 mL of prewarmed mucus removal solution to the tissue.
7. Press and shake the tissue for another 10 min.
Note: Steps H6 and 7 can be repeated if necessary. The time of incubation will depend on the specimen, the mucus, and feces amount. In contrast, if the tissue is rather clean before starting the mucus removal step, one wash step might be sufficient.
Critical: Excessive incubation with mucus removal solution will lead to villi loss. Therefore, the tissue should be washed until the mucus is eliminated (Figure 4). If villi are observed detached in solution, the incubation must be stopped, and the tissue must be immediately transferred to cRPMI (step H8).
Figure 4. Incubation with mucus removal media. (A) An ileal piece of 10 cm open longitudinally previous to mucus removal treatment. (B) Ileal piece after incubation with a mucus removal solution.
8. Add 100 mL of prewarmed cRPMI to the tissue.
9. Press and shake for 10 min.
10. After sufficient mucus removal, transfer the clean tissue to a labeled box containing 150 mL of fresh Custodiol and proceed with the following processing step.
I. Biopsy collection and cryopreservation
This section describes the biopsy collection for RNA, protein isolation, cell isolation, and cryopreservation.
Note: The top layer of the bench protection can be removed to have a clean workspace for the following steps.
1. Place the cooling element on the bench and cover it with a surgical mat or drape.
2. Add Custodiol to your Petri dish and place it on the cooling element.
3. Cut a piece from the proximal side of your tissue piece and transfer it to the Petri dish (Figure 5A).
Figure 5. Example of biopsy tissue collection. (A) Full piece of colon after mucus removal. (B) Example of biopsy collection, where tissue is pinched with tweezers, and 5 mm2 biopsies are taken.
Note: Make sure that you keep the residual tissue submerged and on ice.
Critical: The biopsies should be devoid of any muscle layer and contain only the lamina propria and the mucosal layer. If the tissue is spread, this can be easily achieved by holding the mucosal tissue with the tweezers and cutting it just below. Scissors with curved tips can help to avoid pinching the muscular layer. Biopsy sizes should be approximately 0.5 × 0.5 cm (Figure 5B). Sometimes, it might be easier to first cut a long strip of tissue and then cut it into smaller pieces. If accurate separation of mucosa and submucosa or specific isolation of GALTs is needed, this should be performed under a stereoscope. However, this will delay tissue collection, and redistribution of tasks may be necessary.
Note: The rigidity of the tissue and the characteristics of the mucosal tissue vary along the longitudinal axis of the intestine. While in the small intestine it is easier to cut bigger tissue pieces containing small portions of underlying muscle layer due to the folding, it might be easier to cut small pieces devoid of any muscle layer in the colon.
4. If immune or epithelial cells are to be isolated, start by cutting biopsies for the isolation. Transfer 2 g of tissue or up to 30 pieces of 5 mm2 biopsies to each of the prepared canonical tubes. Continue with section K or M.
5. For protein isolation, cut 8 biopsies and transfer one biopsy to each of the eight tubes prepared for that. Then, transfer them to the box on the dry ice. The box is transferred to liquid nitrogen for long-term storage after all biopsies are collected.
6. For RNA isolation, cut 9 biopsies and transfer three biopsies to each of the three tubes prepared for that. Then, transfer them to the box on the dry ice. The box is transferred to liquid nitrogen for long-term storage after all biopsies are collected.
7. For the collection of samples for biobanking, cut and transfer 10–15 biopsies to each tube and make sure they are submerged in the buffer. Once all the cryovials are filled, transfer the freezing container to 4 °C and incubate it for 1 h. Then, transfer the freezing container to -80 °C. After 24–72 h, transfer the tubes from the freezing container to the white cryo racks prepared in step C1b.
J. Gut disposal
1. Transfer any residual tissue to an autoclaving bag, seal it, and cover it with another autoclaving bag before autoclaving.
2. Dispose of the human tissue in accordance with local regulations.
K. Organoid generation from fresh or cryopreserved tissue
This section describes the generation of region-specific organoids from fresh or cryopreserved tissue. Organoids can be generated directly from fresh tissue (section I) or from thawed cryopreserved tissue (section L). While intact crypt structures and cell clusters can be expected from fresh tissue, crypt isolation from cryopreserved tissue will yield less intact crypt structures and rather cell clusters and single cells.
Critical: The plastic needs to be coated with FBS to avoid any tissue or cells sticking to the plastic.
Note: When generating organoids from cryopreserved tissue, the organoid yield is expected to be lower, and it is more likely that organoids will grow from single cells rather than crypts or cell clusters. Therefore, the time for organoid generation might be 8–12 days instead of 5–7 days. The organoids are then expected to be more sensitive to the dissociation required for expansion. Thus, a low splitting ratio is advised.
1. Organoid line establishment
a. Prepare two 15 mL tubes and one 10 mL pipette per region beforehand by coating them with FBS.
b. Add 10 mL of the dissociation solution to one of the tubes. Keep both tubes on ice.
c. Transfer 15 biopsies to the tube containing the dissociation solution.
d. Incubate it for 30 min (small intestine) or 60 min (large intestine) at 4 °C on a rotating shaker.
e. Remove the dissociation solution.
f. Add 5 mL of ice-cold PBS.
g. Use the coated 10 mL pipette (step K1a) to aspirate and dispense the full volume 20 times.
h. Let the biopsies settle and transfer the supernatant to the FBS-coated 15 mL tube (step K1a).
i. Repeat steps K1f, h, e, g two more times to have a total of 15 mL of cell suspension.
Note: In case of small intestinal cell isolation from fresh tissue, it might be beneficial to use a pre-wet 70 μm strainer to remove villus structures. In order to check if that is necessary, 200 μL of the cell suspension can be added to a microscopy slide and checked under the microscope. If no big structures are visible, this step can be skipped.
i. Centrifuge the suspension at 250× g for 5 min at 4 °C.
Note: A slow break should be used to avoid disturbances to the pellet. In case no pellet has formed, the centrifugation can be repeated.
j. Discard the supernatant and gently resuspend the pellet in a 10 mL of base medium. Repeat the centrifugation step.
k. Resuspend the pellet in the appropriate volume expansion medium containing Y-27632.
Note: The volume added should be equivalent to 2× the pellet size.
l. Mix 10 μL of the cell suspension and 40 μL of Matrigel and seed the full volume to one well of a 24-well plate.
Note: Ideally, 4–6 wells should be plated.
m. Invert the plate and let it polymerize in the cell culture incubator (37 °C, 5% CO2 atmosphere, humidified) for a minimum of 30 min standing on the lid.
Note: By inverting the plate, the settling of cells to the plate bottom is reduced, which improves media diffusion to the organoid and 2D growth.
n. Reinvert the plate to add 500 μL of expansion medium containing Y-27632.
Note: Cell density should not be too low (Figure 6A). A higher cell density (Figure 6B) will support organoid formation (Figure 6C). Dead cells will shrink, whereas viable cells forming organoids present with a sharp cell border (Figure 6D).
o. Change the medium every 2–3 days using expansion medium.
2. Organoid line maintenance and expansion: Once organoids reach 200–300 μm in diameter (Figure 6E, F) they can be expanded.
a. Add 5 mL of cold base medium to a 15 mL tube.
b. Remove the medium from all wells containing organoids without disturbing the Matrigel pellet.
c. Add 1 mL of cold base medium and aspirate and dispense the full volume to release organoids from the Matrigel.
Note: Up to six wells can be pooled in 1 mL of base medium.
d. Transfer the organoid suspension to the 15 mL tube prepared in step K2a.
e. Then, equip a 10 mL pipette with a 200 μL pipette tip and aspirate and dispense the organoid suspension three times.
Note: This step helps to dissolve and dilute the extracellular matrix.
f. Centrifuge the cell suspension at 150× g for 5 min at 4 °C.
g. Remove the supernatant and resuspend the pellet in 50 μL of Accutase per well.
h. Incubate at 37 °C for 45 s.
i. Add 6 mL of cold base medium.
j. Equip a 10 mL pipette with a 200 μL pipette tip and aspirate and dispense the organoid suspension 5–20 times.
Note: The exact number needed depends on the organoid line and needs to be adjusted accordingly. The dissociation should result in a cell suspension containing mostly cell clusters instead of single cells or intact organoids. The success of the dissociation can be checked by microscopy using 10 μL of suspension.
k. Repeat step K2f.
l. Remove the supernatant and resuspend the pellet in a defined volume of expansion medium and Matrigel.
m. Resuspension volume = # wells (pooled for dissociation) × expansion ratio × 45 μL × 0.2 (20% of total volume)
Note: The expansion ratio depends on the density and age of the culture. Directly after isolation, a low expansion should be chosen, e.g., 1:1–1:2, while at later passages, a ratio of 1:6–1:8 can be used. This depends on the individual growth dynamics of each organoid line and needs to be determined.
n. Organoid suspension volume = # wells (desired) × 45 μL × 0.2
o. Matrigel volume = # wells (desired) × 45 μL × 0.8
p. Add 45 μL of the cell suspension Matrigel mix to one well of a 24-well plate.
q. Invert the plate and let it polymerize in the cell culture incubator (37 °C, 5% CO2 atmosphere, humidified) for a minimum of 30 min.
r. Add 500 μL of expansion medium containing Y-27632.
s. Change the medium every 2–3 days with new expansion medium.
t. Repeat step K2a–r every 5–7 days.
Note: The time period between organoid dissociation is dependent on the organoid line as well as the intestinal region. Small intestinal organoids expand slightly slower than colon organoids, which leads to a longer growth period between the dissociations. Organoids can be maintained in culture indefinitely.
Figure 6. Isolation of stem cell–containing crypts and cell suspension for organoid generation. (A) A low and not ideal cell density of single cells in Matrigel dome is shown. (B) A high and optimal cell density of single cells in Matrigel dome is shown. (C) Epithelial cells start to form organoid structures two days after cell isolation. (D) Viable cells and forming organoids show sharp cell boundaries (black arrow), whereas dead cells shrink (white arrow). (E) Organoids derived from fresh descending colon tissue on day four of the fourth passage. (F) Organoids derived from cryopreserved descending colon tissue on day four of the sixth passage. Scale bars are 50 μm (A–D) and 250 μm (E, F). Images A–D show cells isolated from cryopreserved tissue.
L. Thawing of cryopreserved tissue
This section describes how to thaw the cryopreserved tissue biopsies while preserving tissue integrity.
Note: Cryopreservation will reduce the total number of isolated cells 10-fold compared to cell isolation from fresh tissue. However, general population distributions are unaffected, and cells remain functional after cryopreservation. This allows us to study cell isolates at a different time point than the day of tissue processing, to perform sequential analysis of the same region, and to generate autologous co-cultures.
1. Prepare three 50 mL tubes with 20 mL of 0.75 M, 0.375 M, and 0.125 M sucrose and two 50 mL tubes containing 20 mL of 10% FBS/PBS. Keep the tubes at room temperature.
2. Retrieve cryovials containing biopsies from liquid nitrogen and place them on dry ice for transportation.
3. Incubate vials in a water bath at 37 °C until samples are thawed (1–2 min).
Note: Make sure to incubate the biopsies as quickly as possible. Transfer the biopsies before they are fully thawed to avoid cell loss.
4. Pour the content of the vial onto a 100 μm strainer.
5. Collect the biopsies from the strainer and add them to the 50 mL tube containing 20 mL of 0.75 M sucrose.
6. Incubate for 15 min on a horizontal shaker (50–100 rpm) at room temperature.
Note: To minimize cell loss, a low rpm is recommended. Upon thawing, the tissue is fragile. Thus, shear stress should be kept at a minimum while making sure the tissue is in motion.
7. Pour the content of the 50 mL tube onto a 100 μm strainer.
8. Collect the biopsies and add them to the 50 mL tube containing 20 mL of 0.375 M sucrose.
9. Incubate for 15 min on a shaker (50–100 rpm) at room temperature.
10. Pour the content of the 50 mL tube onto a 100 μm strainer.
11. Collect the biopsies and add them to the 50 mL tube containing 20 mL of 0.125 M sucrose.
12. Incubate for 15 min on a shaker (50–100 rpm) at room temperature.
13. Pour the content of the 50 mL tube onto a 100 μm strainer.
14. Collect the biopsies and add them to the 50 mL tube containing 20 mL of 10% FBS/PBS.
15. Incubate for 10 min on a shaker (50–100 rpm) at room temperature.
16. Repeat steps L13–15.
17. Pour the content of the 50 mL tube onto a 100 μm strainer.
18. Collect the biopsies and proceed with desired cell isolation (section K, M–O).
M. Epithelial cell isolation for FACS analysis or epithelial cell removal prior to immune cell isolation
This section describes how to either isolate epithelial cells for downstream analysis or remove epithelial cells from the tissue biopsies to allow for purer immune cell isolation.
Note: Starting material can be freshly taken (section I) or thawed cryopreserved biopsies (section L).
Note: If a co-culture is the aim of immune cell isolation, the addition of primocin (100 μg/mL) to epithelial cell removal media is recommended to avoid possible contaminations at later stages.
1. Prepare tubes:
a. Tube 1: 50 mL tube with 25 mL of epithelial cell removal medium.
b. Tube 2: 50 mL tube with 25 mL of epithelial cell removal medium.
c. Tube 3: 50 mL tube with 25 mL of cRPMI.
d. Tube 4: Empty collection 50 mL tube labeled as an epithelial fraction.
2. Prewarm tubes 1, 2, and 3 (steps M1a–c).
3. Place tube 4 on ice (step M1d).
4. Transfer the biopsies to tube 1 (step M1a) containing 25 mL of epithelial removal medium.
5. Incubate for 20 min at 37 °C on a horizontal shaker (350 rpm).
Note: In this step, the sheer stress should be strong to allow epithelial cell detachment.
6. Vortex the tube with biopsies vigorously for 30 s.
7. Pre-wet a 70 μm strainer with 5 mL of cRPMI.
8. Pour the content of tube 1 onto the 70 μm strainer and collect the flowthrough in tube 4 (step M1d) placed on ice.
9. Collect the biopsies from the strainer and add them to tube 2 (step M1b) containing 25 mL of epithelial cell removal medium.
10. Incubate for 20 min at 37 °C on a shaker (350 rpm).
11. Pour the content onto the 70 μm strainer and collect the flowthrough in tube 4.
12. Collect the biopsies from the strainer and add them to tube 3 (step M1c) containing 25 mL of cRPMI.
13. Discard the flowthrough (enriched on fibroblasts), transfer the biopsies for immune cell isolation (section N), or discard them if the interest is in the epithelial fraction.
14. To recover the epithelial cells, centrifuge tube 4 at 400× g for 10 min at 4 °C.
15. Discard the supernatant and resuspend the pellet in 1 mL of cRPMI.
Note: Centrifugation of tube 4 and resuspension on cRPMI can be done after the first collection (step M6) or at the end (step M9).
Note: Two incubations with epithelial cell removal media are the most efficient way to remove epithelial cells. While further incubations will remove more epithelial cells, it is not time efficient. However, further incubations can be done if the purpose is to obtain high amounts of epithelial cells.
Note: The epithelial fraction can then be used for downstream experiments, such as FACS characterization, sorting, or RNA-seq. For organoid generation, epithelial cells as single-cell suspension should be processed immediately, and depending on the assay, the addition of Y-27632 should be considered.
Note: If done with fresh biopsies, cells from the epithelial fraction can also be frozen for later use by resuspending 106–107 cells in 1 mL of CryoStor buffer.
N. Immune cell isolation for FACS analysis or organoid-immune cell co-culture establishment
This section describes three different digestion methods for immune cell isolation that support the isolation of different immune cell populations (see Validation).
Note: For immune cell isolation, the biopsies from section M are used. Even if the focus is on immune cells, the epithelial cell isolation step (section M) should be performed to reduce the number of contaminating epithelial cells and allow for better tissue digestion.
1. Prewarm 5 mL of cRPMI per gram of tissue to be digested at 37 °C.
2. Fill and prewarm a Duran crystallizing beaker with water on a magnetic stirrer at 37 °C.
Note: Depending on the enzymes used, specific immune cell populations will be enriched (see Validation). Since there is a wide range of downstream applications, factors such as cell number, enrichment, relative amount of immune cell populations, or contamination with epithelial cells may be considered to get the optimal required output (see Validation). If only a general overview of general immune cell populations is desired, it is recommended that enzymatic mixture B (collagenase IV and DNaseI) is used.
Note: For enzymatic mixtures A and B, 35 mL tubes are used, whereas for enzymatic mixture C, c-tubes are required.
Note: Be aware that depending on the collagenase blend, specific cell receptors can be cleaved, and this should be taken into account when designing the experiment [2].
3. Weigh the biopsies from section M and add 0.5–1 g of tissue into a 35 mL tube containing 5 mL of prewarmed cRPMI (Figure 7A).
Figure 7. Example of biopsy handling for immune cell isolation. (A) Biopsies before (left) and after (right) being cut into tiny fragments with scissors. (B) Result of biopsy digestion after incubation with enzymatic mixtures.
4. Cut the tissue into tiny fragments using scissors (Figure 7A).
5. Add the enzymes to the corresponding tube (see Recipes 21–23).
Note: Fresh preparation of these enzymes is recommended; stock dilutions with higher concentrations can be prepared beforehand and stored at -20 °C in small aliquots to avoid refreezing or enzyme loss.
6. For enzymatic mixtures A and B, add a magnetic stirrer into each tube.
7. Transfer the tubes to the water bath prepared in step N2 and incubate them for 45 min under constant agitation at 350 rpm.
Note: Using different-sized magnetic stirrers will affect cell recovery. It is important to use magnetic stirrers of the same size (15 mm length and 6 mm diameter) and ensure that there are no ridges to avoid cell loss.
8. For enzymatic mixture C, place the c-tube in the gentleMACS Octo Dissociator and run protocol MultiB for 1 h at 37 °C.
Note: From this step onward, the samples are processed in the same way, independently of the enzyme mixture used.
9. Pre-wet a 40 μm strainer with 5 mL of cRPMI.
Note: Other strainers, such as 70 or 100 μm, can be used. However, in our experience, large-size strainers do not increase immune cell recovery; instead, they allow the recovery of more epithelial and dead cells.
10. After incubation, pour the content of the tube onto the pre-wetted 40 μm strainer into a fresh collection tube.
11. Wash tube and magnetic stirrer with 5 mL of cRPMI and transfer the full volume onto the 40 μm strainer.
12. Wash the 40 μm strainer with another 5 mL of cRPMI.
13. Collect both washes in the same tube as the isolated cells.
Note: Isolated cells will be in the flowthrough.
14. Centrifuge the cells at 400× g for 10 min at 4 °C.
15. Discard the supernatant and resuspend the pellet containing the isolated immune cells in 1 mL of cRPMI.
16. Keep it on ice and use it for characterization, co-culture, or any other application.
Note: Ensure that the digestion is complete (Figure 7B). If digestion is not complete (tissue pieces are still visible), place the undigested tissue and repeat the protocol starting at step N5. This will increase the time for isolation and the final number of total isolated cells. If the tissue is not completely digested even after the second digestion, the biopsies will not be sufficiently minced, or the muscular layer will be carried over when taking the biopsy. Alternative methods to facilitate complete digestion can also be applied, such as vigorously pipetting the solution regularly or using a homogenizer to disrupt the tissue.
O. Co-culture of organoids and lamina propria immune cells
Note: To generate a co-culture of organoids and lamina propria immune cells, organoid generation needs to be performed in advance, which takes approximately 4 weeks. Organoids should be expanded at least two times or until the desired number of wells is reached. Immune cells are isolated from fresh (section I + section M and N) or cryopreserved tissue (section L + section M and N) on the day of the co-culture experiment. If an autologous co-culture is desired, immune cells must be isolated from cryopreserved tissue of the same donor as used for organoid generation.
1. Organoid culture
Note: Grow organoids from fresh or cryopreserved tissue as described above (section K). Passage organoids at least 2–3 times before using them for co-culture establishment.
a. Three days prior to the planned immune cell isolation, seed the desired number of wells for co-culture and let the organoids grow for 3 days in a growth medium (see step K2 for further details).
Note: On day 3 of the culture, the organoids are harvested and co-cultured with freshly isolated immune cells (see step O2).
2. Immune cell enrichment with gradient centrifugation
Note: This step is critical to remove unwanted cell types such as fibroblasts, epithelial cells, and dead cells while enriching for total lamina propria immune cells. Specific immune cell types may be isolated depending on the individual needs using FACS or MACS.
Note: This step is performed directly after immune cell isolation, as described in section N. The addition of primocin (100 μg/mL) to epithelial cell removal media is recommended to avoid possible contaminations in co-culture.
Note: Thaw Matrigel on ice. Ensure Percoll buffers and centrifuge are at room temperature.
a. Centrifuge isolated cells from step N16 in a 15 mL tube at 400× g for 5 min at room temperature.
b. Prepare a new 15 mL tube filled with 5 mL of 80% Percoll.
c. Resuspend the cell pellet in 1 mL of 40% Percoll.
d. Then, add 9 mL of 40% Percoll to a total volume of 10 mL.
e. Mix the cell suspension by slowly pipetting up and down three times using a 10 mL pipette.
f. Transfer the cell suspension with a 10 mL pipette to a tube containing 5 mL of 80% Percoll prepared in step O2b.
Critical: To avoid mixing the two density layers, add the cell suspension dropwise on top of the 80% Percoll layer while holding the tube at a 45° angle. This will minimize the mixing of the layers and ensure sufficient gradient centrifugation.
Critical: Slow pipetting of the cell suspension is critical to avoid mixing of both gradients. Mixing of the gradients will result in reduced numbers of immune cells.
g. Immediately centrifuge the cell suspension at 1,360× g for 20 min at room temperature with reduced acceleration and without a break (acceleration = 2, deceleration = 0).
Critical: Excessive acceleration or abrupt stopping of the centrifuge will mix the gradients and result in cell loss.
h. Carefully remove the tube from the centrifuge and check successful separation.
Note: A white immune cell layer will be visible between the two layers at the 5 mL position. Depending on the cell number, the layer might be more transparent or not clearly visible.
i. Remove the upper Percoll layer without harming the immune cell layer.
j. Carefully take up the white immune cell layer at the 5 mL position using a P1000 pipette.
Note: Take up as little as possible Percoll, as it may reduce the overall cell yield and may increase the risk of contamination of other cell types. In case the immune cell layer is not clearly visible, take up 1 mL of Percoll at a 5 mL position.
k. Transfer the cells to a 50 mL tube and add 29 mL of PBS-/-.
l. Slowly invert the tube three times and centrifuge at 400× g for 10 min at 4 °C.
Note: From now on, no specific break is required (acceleration = 9, deceleration = 9) during centrifugation.
m Discard the supernatant and resuspend the pellet in 1 mL of PBS-/-.
n. Repeat the wash step (steps O2k–m).
o. Use 10 μL of cell suspension to count the cells.
p. Distribute the volume to 15 mL tubes so that each has 0.6 × 106 cells.
Note: Each tube contains the immune cells that will be combined with organoids from one well of a 24-well plate. Depending on your experimental design and the immune cells yield, you can either set up several wells containing co-cultures, or you can include controls containing only immune cells.
q. Centrifuge the cells at 400× g for 10 min at 4 °C.
r. Discard the supernatant and keep cells on ice until mixing with organoids.
3. Organoid harvest
a. On day 3 of culture, the organoids are harvested by adding 1 mL of ice-cold base medium to one well containing organoids. Pipette up and down to release the organoids from the Matrigel.
b. Transfer the organoids of one well of a 24-well plate to a 15 mL tube filled with 5 mL of base medium.
c. Remove Matrigel by pipetting up and down three times with a 10 mL pipette equipped with a 200 μL pipette tip.
d. Centrifuge the organoid suspension at 150× g for 5 min at 4 °C.
e. Remove the supernatant and place it on ice.
4. Co-culture setup
a. Remove any remaining supernatant of the immune cells (step 2r) and organoid pellet (step 3e) placed on ice before.
b. Thoroughly resuspend the immune cells (0.6 × 106 cells) in 50 μL of ice-cold Matrigel by pipetting 10 times up and down.
Critical: For immune cell movement and migration, it is detrimental to resuspend them in an extracellular matrix.
c. Thoroughly resuspend the organoid pellet using 45 μL of immune cells in Matrigel by pipetting up and down three times while pushing the tip toward the tube bottom.
d. Distribute the 45 μL Matrigel containing immune cells and organoids into three drops in one well of a 24-well plate. The density of the organoids and immune cells is shown in Figure 8A.
Note: Depending on the reading, the plating of the co-culture may be adjusted, e.g., seeding into a flat dome.
e. Transfer the plate to the incubator (37 °C, 5% CO2 atmosphere, humidified) and let it polymerize for 1 h.
f. Then, add 500 μL of growth medium and place it back in the cell culture incubator.
5. Live imaging of co-culture
Note: This step is used to monitor how the immune cells move and to identify how the cells interact with the organoids, as shown in Figure 8B.
a. Prewarm the stage incubator to 37 °C and 5% CO2.
b. Place the co-culture plate on the microscope and focus on the area of interest.
c. Acquire images every 1–5 min for at least 30 min.
6. Fixation of co-culture
a. Carefully remove the medium by tilting the plate without touching the Matrigel dome.
Note: Medium can be sampled to measure cytokine release.
b. Add 500 μL of co-culture fixative to the plate and incubate it for 20 min at room temperature.
Critical: To maintain the 3D structure of the Matrigel dome, the fixative needs to contain 0.1% glutaraldehyde.
c. Remove the fixative and wash two times with 500 μL of PBS-/- for 5 min.
d. Add PBS-/- and proceed with downstream experiments.
Note: Fixed co-culture can be stored up to 24 h at 4 °C until embedding or whole-mount staining.
Figure 8. Representative brightfield images of lamina propria–derived immune cells co-cultured with intestinal organoids. (A) Expected cell and organoid density when following the protocol. Scale bar: 100 μm. (B) Interaction of immune cells with the organoids, which can be expected to be observed when performing live cell imaging. Scale bar: 25 μm.
Data analysis
Statistical tests
GraphPad Prism v10.0.0 was used for plotting and statistical calculation. 2-way ANOVA and Tukey multiple comparison tests were performed to test for significance. Significance was achieved when p-value < 0.05.
Validation of protocol
A. Comparison of different immune cell isolation protocols
In the following section, the three different enzymatic digestion methods were compared using tissue from three different patients. Cryopreserved tissue from mid jejunum was analyzed by flow cytometry using a general panel for immune cells and epithelial identification (Table 1).
Table 1. Panel used for immunostaining and flow cytometry analysis
Marker Label cell of interest Company Clone Conjugate Cat # Volume (μL)
CD45 All immune cells BD Biosciences HI30 BV510 563204 5
7AAD Viability BioLegend 420404 1
EPCAM Epithelial cells BioLegend 9C4 PE 324206 1.5
CD3 T cells BD Biosciences UCTH1 BUV496 612940 5
CD19 B cells BioLegend HIB29 BV750 302262 5
HLA-DR Antigen-presenting cells BD Biosciences G46-6 BUV395 564040 1
CD14 Macrophages BD Biosciences M5E2 BUV737 612763 2.5
CD11c Dendritic cells BD Biosciences B-Ly6 BB515 564490 1
CD56 NK cells BioLegend HCD56 APC/Cy7 318332 5
CD103 Intra-epithelial lymphocytes BioLegend Ber-ACT8 APC 350216 2.5
Figures 9A and 9B show that extraction can vary between human samples. However, if the interest is in epithelial cells, collecting the epithelial fraction isolated with the removal media (section M) is the most efficient method. If immune cells are the focus, then enzymatic mixtures are the best approach, especially enzymatic mixture B (ColIV + DNaseI), which manages the isolation of approximately 5 million immune cells per gram of tissue. Data also show that each enzymatic mixture enriches different immune cell subtypes; while enzymatic mixture B (ColIV + DNaseI) allows for a higher retrieval of general immune cells as well as enrichment of B cells (Figure 9A, C), the enzymatic mixture A (Liberase + DNaseI) is best if NK cells are the object of the study (Figure 9D). Both enzymatic mixtures A and B show similar ability to obtain T cells, dendritic cells, and macrophages (Figure 9E–G). In contrast, enzymatic mixture C (MTKD1) leads to a high overall immune cell loss (Figure 9A). However, immune cells are present in a higher proportion (Figure 9H), which could be useful for certain applications such as bulk RNAseq.
Figure 9. Comparison of different enzymatic mixtures for cell isolation using cryopreserved jejunal samples from three different human donors. (A) Total number of immune cells isolated per gram of tissue. (B) Total number of epithelial cells isolated per gram of tissue. (C) Number of B cells isolated per gram of tissue. (D) Number of NK cells per gram of tissue. (E) Number of T cells per gram of tissue. (F) Number of dendritic cells per gram of tissue. (G) Number of macrophages per gram of tissue. (H) Immune cell enrichment normalized to the total amount of isolated cells. *p < 0.05.
B. Validation in other published research articles
This protocol or parts of it have been used and validated in the following research articles:
Hensel et al. [13]. SLE serum induces altered goblet cell differentiation and leakiness in human intestinal organoids. EMBO Molecular Medicine.
Hensel et al. [14]. Protocol for generating and analyzing organ-on-chip using human and mouse intestinal organoids. STAR Protocols.
Supplementary information
The following supporting information can be downloaded here:
1. Excel file for biobank
Acknowledgments
We would like to thank Hugo de Jonge for providing the R-Spondin 1 cell line. We also thank Dr. Stephanie Muenchau for her help setting up the experiment and their input during the organoid generation from frozen biopsies. We also express our gratitude to Thomas Rückle and Christian Tidona for their invaluable help. Special recognition goes to the tissue donors, the surgeons, Novabiosis, the DTI Foundation, and the I3PT Biobank, who made this study possible. We specifically thank Joaquim Albiol, Fernando Mosteiro, and his team for their dedication. We thank Richard Fairless (Department of Neurology at the University Heidelberg) and specifically Katharina Schmitz for her help with tissue processing and Stefan Fritz (Dr. Franz Köhler Chemie) for the generous donation of Custodiol for research purposes. Christoph Becker and Daigen Xu's critical feedback and guidance significantly improved the quality of our work. This research was funded by Merck KGaA, Darmstadt, Germany (M.R.D., I.V.H., M.S., I.M.S., B.S., S.D., and S.D.) and by Johnson & Johnson Innovative Medicine, USA (K.K., E.G.E., O.M., and E.S.). This protocol was adapted and modified from Hensel et al. [13,14].
Competing interests
The authors declare no conflict of interest.
Ethical considerations
Human whole intestinal tissue samples were procured from donors via the following: 1) Novabiosis, Inc. (Research Triangle Park, Durham, North Carolina, USA) following ethical committee approval from the Organ Procurement Organizations (OPO), in line with the consent and deidentification guidelines established by the OPOs and the United Network for Organ Sharing (UNOS), under the US transplantation network framework. Immediate family members of the donors granted permission for organ donation while preserving the donor’s privacy. This donation approval is according to the guidelines provided by the federal organization UNOS and the Federal Drug Administration (FDA). 2) I3PT Biobank and the DTI Foundation (Barcelona, Spain). The tissue donations were processed following standard operating procedures with the appropriate approval of the Ethics and Scientific Committees.
All research procedures were conducted adhering to the principles specified in the WMA Declaration of Helsinki.
References
Bujko, A., Atlasy, N., Landsverk, O. J., Richter, L., Yaqub, S., Horneland, R., Øyen, O., Aandahl, E. M., Aabakken, L., Stunnenberg, H. G., et al. (2018). Transcriptional and functional profiling defines human small intestinal macrophage subsets. J Exp Med. 215(2): 441–458.
Doyle, C. M., Fewings, N. L., Ctercteko, G., Byrne, S. N., Harman, A. N. and Bertram, K. M. (2022). OMIP 082: A 25‐color phenotyping to define human innate lymphoid cells, natural killer cells, mucosal‐associated invariant T cells, and γδ T cells from freshly isolated human intestinal tissue. Cytometry Part A. 101(3): 196–202.
Uronen-Hansson, H., Persson, E., Nilsson, P. and Agace, W. (2014). Isolation of Cells from Human Intestinal Tissue. Bio Protoc. 4(7): e1092.
Hughes, S. M., Shu, Z., Levy, C. N., Ferre, A. L., Hartig, H., Fang, C., Lentz, G., Fialkow, M., Kirby, A. C., Adams Waldorf, K. M., et al. (2016). Cryopreservation of Human Mucosal Leukocytes. PLoS One. 11(5): e0156293.
Hughes, S. M., Ferre, A. L., Yandura, S. E., Shetler, C., Baker, C. A. R., Calienes, F., Levy, C. N., Astronomo, R. D., Shu, Z., Lentz, G. M., et al. (2018). Cryopreservation of human mucosal tissues. PLoS One. 13(7): e0200653.
Urbano, P. C. M., Angus, H. C. K., Gadeock, S., Schultz, M. and Kemp, R. A. (2022). Assessment of source material for human intestinal organoid culture for research and clinical use. BMC Res Notes. 15(1): 1–8.
Campbell, J., Berry, J. and Liang, Y. (2019). Anatomy and Physiology of the Small Intestine. In: Shackelford's Surgery of the Alimentary Tract, 2 Volume Set (8th Edition, pp. 817–841). Elsevier Inc.
Ma, Z. F. and Lee, Y. Y. (2020). Small intestine anatomy and physiology. In: Clinical and Basic Neurogastroenterology and Motility. (pp. 101–111). Elsevier Inc.
Phillips, M., Patel, A., Meredith, P., Will, O. and Brassett, C. (2015). Segmental colonic length and mobility. Ann R Coll Surg Engl. 97(6): 439–444.
Rubin, D. C. and Shaker, A. (2009). Small Intestine: Anatomy and Structural Anomalies. In: Podolosky, D., Camilleri, M., Fitz, J., Kalloo, A., Shanahan, F. and Wang, T. (Eds.). Textbook of Gastroenterology (6th Edition, Vol. 1, pp. 1085–1107). Wiley.
Treuting, P. M., Valasek, M. A. and Dintzis, S. M. (2012). Upper Gastrointestinal Tract. In: Comparative Anatomy and Histology (1st Edition, Issue Figure 5, pp. 155–175). Elsevier Inc.
Umanskiy, K. and Matthews, J. (2008). Colon: Anatomy and Structural Anomalies. In: Podolosky, D., Camilleri, M., Fitz, J., Kalloo, A., Shanahan, F. and Wang, T. (Eds.). Textbook of Gastroenterology (6th Edition, Vol. 1, pp. 1369–1385). Wiley.
Hensel, I. V., Éliás, S., Steinhauer, M., Stoll, B., Benfatto, S., Merkt, W., Krienke, S., Lorenz, H. M., Haas, J., Wildemann, B., et al. (2024). SLE serum induces altered goblet cell differentiation and leakiness in human intestinal organoids. EMBO Mol Med. 16(3): 547–574.
Hensel, I. V., Steinhauer, M., Fairless, R. and Resnik-Docampo, M. (2024). Protocol for generating and analyzing organ-on-chip using human and mouse intestinal organoids. STAR Protoc. 5(2): 103037.
Article Information
Publication history
Received: Jun 20, 2024
Accepted: Oct 28, 2024
Available online: Nov 20, 2024
Published: Jan 5, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Utilizing FRET-based Biosensors to Measure Cellular Phosphate Levels in Mycorrhizal Roots of Brachypodium distachyon
SZ Shiqi Zhang
LJ Lucas Jurgensen
MH Maria J. Harrison
Published: Vol 15, Iss 2, Jan 20, 2025
DOI: 10.21769/BioProtoc.5158 Views: 399
Reviewed by: Demosthenis ChronisJuliane K Ishida Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in New Phytologist Jun 2022
Abstract
Arbuscular mycorrhizal (AM) fungi engage in symbiotic relationships with plants, influencing their phosphate (Pi) uptake pathways, metabolism, and root cell physiology. Despite the significant role of Pi, its distribution and response dynamics in mycorrhizal roots remain largely unexplored. While traditional techniques for Pi measurement have shed some light on this, real-time cellular-level monitoring has been a challenge. With the evolution of quantitative imaging with confocal microscopy, particularly the use of genetically encoded fluorescent sensors, live imaging of intracellular Pi concentrations is now achievable. Among these sensors, fluorescence resonance energy transfer (FRET)-based biosensors stand out for their accuracy. In this study, we employ the Pi-specific biosensor (cpFLIPPi-5.3m) targeted to the cytosol or plastids of Brachypodium distachyon plants, enabling us to monitor intracellular Pi dynamics during AM symbiosis. A complementary control sensor, cpFLIPPi-Null, is introduced to monitor non-Pi-specific changes. Leveraging a semi-automated ImageJ macro for sensitized FRET analysis, this method provides a precise and efficient way to determine relative intracellular Pi levels at the level of individual cells or organelles.
Key features
• This protocol describes the use of FRET biosensors for in vivo visualization of spatiotemporal phosphate levels with cellular and subcellular resolution in Brachypodium distachyon.
• An optimized growth system can allow tracing of Pi transfer between AM fungi and host root.
Keywords: Arbuscules Fluorescence resonance energy transfer (FRET) Sensitized FRET Live imaging Symbiosis Roots Phosphate
Background
Arbuscular mycorrhizal (AM) fungi establish one of the most prevalent mutualistic relationships with plants, primarily delivering phosphorus in the form of phosphate (Pi) to their hosts for carbon in return. The symbiosis is characterized by AM fungal hyphae penetrating the root epidermis, progressing through the root to the cortex, and forming arbuscules within cortical cells. The arbuscules, intricate tree-like structures, serve as the primary sites for nutrient exchange.
Mycorrhizal roots contain a diverse array of cortical cell conditions including cells hosting intracellular hyphae and cells containing arbuscules at different stages of development, from initiation to maturity and eventual collapse [1,2]. This diversity in arbuscule development across the cortical cells is predicted to result in variation in Pi transfer, as periarbuscular membrane-resident Pi transporters show arbuscule stage–specific expression [3–6]. Therefore, variance in Pi content in these cells is anticipated. In addition, plastids in certain colonized cells become highly stromulated [7,8], suggesting increased metabolic activity that may result in intracellular shifts in Pi levels.
Furthermore, the process of Pi uptake in mycorrhizal roots markedly contrasts with that of non-mycorrhizal roots. While non-mycorrhizal roots absorb Pi through their epidermal cells, transporting it apoplastically and/or sequentially to the cortex, endodermis, and vasculature, mycorrhizal roots receive Pi uptake directly into their inner cortical cells before moving it to the endodermis and vasculature [5,6]. Epidermal Pi transporter gene expression is downregulated in mycorrhizal roots, suggesting that the epidermis in mycorrhizal roots plays a less significant role in Pi absorption [9–11]. Given these distinct pathways for Pi uptake, investigating the cellular Pi response dynamics within mycorrhizal roots is likely to provide new insights into root Pi distribution and homeostasis.
Genetically encoded fluorescent sensors have emerged as powerful tools for monitoring analytes with cellular resolution [12,13]. These fluorescent sensors can be introduced into plants and expressed from constitutive or tissue/cell-type-specific promoters. They have the ability to bind reversibly to their analytes, translating their abundance into fluorescence changes. This facilitates real-time monitoring of molecular changes at intracellular or subcellular levels. Biosensors, targeting ions like Ca2+, H+, and Zn2+, as well as hormones such as ABA and auxin, and monitoring protein activities such as nitrate transporters, have been created and used in plants [12]. Their application has advanced knowledge of ion and hormone dynamics during plant development and in response to abiotic stress [13–15].
Among the various biosensors, fluorescence resonance energy transfer (FRET)-based sensors have proven especially reliable [12,13]. FRET occurs when a donor and an acceptor fluorescent molecule are in close proximity, and there is spectral overlap between the donor emission and the acceptor excitation; energy then transfers from the excited donor to the acceptor, quenching the donor’s fluorescence and potentially enhancing fluorescence of the acceptor. FRET biosensors generally feature a ligand-binding domain attached to two green fluorescent protein (GFP) spectral variants, typically cyan (CFP) and yellow (YFP). When the ligand binds, it triggers a conformational change that alters the proximity of the donor and acceptor and affects the FRET efficiency. This change is observed as a shift in the intensity ratio of the two fluorescent proteins, termed the FRET ratio [13]. The observed ratio directly corresponds to the ligand concentration within a range defined by the binding affinity and, ultimately, by the sensor's saturation point [16–18]. Furthermore, FRET-sensitized emission is a valuable method to measure FRET; adjusting for issues like donor spectral bleed-through and acceptor cross-excitation allows a precise determination of FRET-derived acceptor fluorescence [19–21]. Such adjustments are vital when analyzing ligand concentrations across various cell types or comparing data from different samples to pinpoint gene or protein roles.
To measure Pi content in mycorrhizal roots, we introduced a Pi biosensor, cpFLIPPi-5.3m, into Brachypodium distachyon plants and targeted it to the cytosol or plastid [22]. In each case, we included a control sensor, cpFLIPPi-Null, which has a mutation that prevents Pi binding. These sensors had previously been optimized and used successfully to monitor intracellular Pi content in Arabidopsis thaliana [23,24]. The authors had shown that the control sensor is Pi-independent and any FRET ratio shifts reflect non-Pi specific changes, such as intracellular ionic shifts. Thus, when the control sensor exhibits no FRET ratio shift, it suggests that the observed changes in the Pi sensor are genuinely due to intracellular Pi fluctuations. This study outlines our method of employing these sensors to track Pi dynamics during AM symbiosis and our use of a semi-automated ImageJ macro to efficiently extract quantitative data for sensitized FRET analysis.
Materials and reagents
Biological materials
1. All Brachypodium distachyon (line Bd21-3) transgenic lines used in this protocol can be obtained from Maria Harrison’s lab (under a Boyce Thompson Institute Material Transfer Agreement). The lines are grouped into categories as follows:
a. For the quantification of cytosolic Pi levels:
Transgenic lines expressing the sensor and its controls from a mycorrhiza-inducible, cell-type-specific promoter, BdPT7. Fluorescent signals from these lines are only from cells containing arbuscules.
BdPT7::cpFLIPPi-5.3m
BdPT7::cpFLIPPi-Null
BdPT7::eCFP
BdPT7::cpVenus
WT (used as a negative control to remove the potential background during image analysis)
All lines are needed for Pi imaging for each experiment.
Transgenic lines expressing the sensor and controls from the constitutive promoter ZmUb1 promoter. Fluorescent signals from these lines occur in all cell types. We failed to generate a ZmUb1::cpVenus line so the BdPT7::cpVenus is used as one of the controls for ZmUb1::cpFLIPPi-5.3m.
ZmUb1::cpFLIPPi-5.3m
ZmUb1::cpFLIPPi-Null
ZmUb1::eCFP
BdPT7::cpVenus
WT (used as a negative control to remove the potential background during image analysis)
b. For the quantification of plastidic Pi levels:
Transgenic lines expressing a plastid-targeted sensor and its controls from the BdPT7 promoter. Fluorescent signals from these lines occur in the plastid only in cells containing arbuscules.
BdPT7::plastid-cpFLIPPi-5.3m
BdPT7::plastid-cpFLIPPi-Null
BdPT7::plastid-eCFP
BdPT7::plastid-cpVenus
WT (used as a negative control to remove the potential background during image analysis)
2. AM fungal spores. Diversispora epigaea is used in the protocol as an example. It can be obtained from the International Collection of (Vesicular) Arbuscular Mycorrhizal Fungi (INVAM). Other species of AM fungi are also available from this collection. We have also used these lines successfully with Rhizophagus irregularis. A commercial vendor, Premier Tech (Canada), can supply Rhizophagus irregularis (ready-for-use spores).
Reagents
1. Bleach (Pure Bright, containing 5%–7% sodium hypochlorite)
2. Sterile deionized water
3. Agar (Millipore-Sigma, catalog number: A7921)
4. Benomyl [Methyl 1-(butylcarbamoyl)-2-benzimidazolecarbamate] (Millipore-Sigma, catalog number: 381586)
5. Ca(NO3)2·4H2O (calcium nitrate tetrahydrate) (Millipore-Sigma, catalog number: C2786-500G)
6. KNO3 (potassium nitrate) (Millipore-Sigma, catalog number: 221295-100G)
7. MgSO4·7H2O (magnesium sulfate heptahydrate) (Millipore-Sigma, catalog number: 221295-100G)
8. NaFeEDTA [ethylenediaminetetraacetic acid iron(III) sodium salt] (Millipore-Sigma, catalog number: EDFS-100G)
9. KH2PO4 (potassium phosphate monobasic) (Millipore-Sigma, catalog number: P0662-25G)
10. H3BO3 (boric acid) (Millipore-Sigma, catalog number: B0394-100G)
11. Na2MoO4·2H2O (sodium molybdate dihydrate) (Millipore-Sigma, catalog number: 331058-5G)
12. ZnSO4·7H2O (zinc sulfate heptahydrate) (Millipore-Sigma, catalog number: 221376-100G)
13. MnCl2·4H2O [manganese(II) chloride tetrahydrate] (Millipore-Sigma, catalog number: 221279-100G)
14. CuSO4·5H2O [copper(II) sulfate pentahydrate] (Millipore-Sigma, catalog number: 209198-5G)
15. CoCl2·6H2O [cobalt(II) chloride hexahydrate] (Millipore-Sigma, catalog number: 255599-5G)
16. HCl (hydrochloric acid) (Millipore-Sigma, catalog number: 258148-25ML)
17. MES (C6H13NO4S) (Millipore-Sigma, catalog number: M3671-50G)
18. NaOH (sodium hydroxide) (Millipore-Sigma, catalog number: 221465-500G)
Solutions
1. Modified 1/4 strength Hoagland solution with 20 μM potassium phosphate (see Recipes)
Recipes
1. Modified 1/4 strength Hoagland solution with 20 μM potassium phosphate
Reagent Final concentration Note
Ca(NO3)2·4H2O 1.25 mM
KNO3 1.25 mM
MgSO4·7H2O 0.5 mM
NaFeEDTA 0.025 mM
KH2PO4 0.02 mM The phosphate levels can be varied by adjusting the amount of this reagent.
H3BO3 5 μM
Na2MoO4·2H2O 0.12 μM
ZnSO4·7H2O 0.5 μM
MnCl2·4H2O 1 μM
CuSO4·5H2O 0.25 μM
CoCl2·6H2O 0.19 μM
HCl 6.25 μM
MES (C6H13NO4S) 0.25 mM
Adjust the final pH to 6.1 using NaOH.
Laboratory supplies
1. 14 mL culture polypropylene tubes (MTC-Bio, catalog number: UX-34501-06)
2. Regular-weight seed germination paper (Anchor Paper Co, catalog number: SD3815L)
3. Sterile Petri dish (100 × 10, Carolina, catalog number: 741248; 100 × 20 mm, Carolina, catalog number: 741252; a large one, approximately 14 cm in diameter, used for washing substrate from roots)
4. Parafilm (Sigma-Aldrich, catalog number: P7543)
5. 20 cm plastic cones (Stuewe & Sons, Inc., catalog number: SC10U)
6. 9 cm (3-1/2”) diameter plastic pot (Flinn Scientific, catalog number: FB0652)
7. Humidity domes (Global Industrial, catalog number: CK64081)
8. Growth substrate (for example, sand, gravel, or turface, which can be purchased from a general supplier, e.g., Lowes or Home Depot)
9. MF Millipore membrane filter, 0.45 μm pore size (Millipore-Sigma, catalog number: HAWP9000)
10. Microscope slides (ideally 75 mm × 25 mm, Carolina, catalog number: 632010) and coverslips (ideally 20 mm × 20 mm, Carolina, catalog number: 633009)
11. Aluminum foil
12. Fine tip tweezers (Fisherbrand, catalog number: 12-000-122)
13. A digital camera or other device that can take photos, such as a personal cell phone
14. Spray bottles (UNLINE, catalog number: S-11686)
Equipment
1. Class I laminar flow hood (e.g., Thermo Scientific HeraguardTM ECO Clean Bench, catalog number: 51029701)
2. Growth chambers or greenhouses to grow plants (e.g., Conviron reach-in growth chamber, model: PGR15)
3. Stereomicroscope (Olympus, model: SZX-12)
4. Upright confocal microscope (Leica, model: SP5), see General note 1 for details
Software and datasets
1. Fiji (ImageJ, free, no license needed, can be downloaded via the link: https://imagej.net/downloads)
The sensitized FRET analysis Macro can be copied via the link: https://nph.onlinelibrary.wiley.com/action/downloadSupplement?doi=10.1111%2Fnph.18081&file=nph18081-sup-0001-SupInfo.pdf)
Procedure
A. Plant material preparation
1. AM fungal spore preparation (optional if inoculating with AM fungi)
The preparation of AM fungal spores varies among fungal species and research purposes. If you have an established method, follow it. Otherwise, consider these options:
a. Obtain ready-for-use spores from the International Collection of (Vesicular) Arbuscular Mycorrhizal Fungi (INVAM) or commercial vendors like Premier Tech, Canada.
b. Refer to spore preparation methods in [26] or [27].
c. In cases when specific fungal species are not required, directly inoculate plants with commercial or home-made mixed propagule inoculum as described in these studies [28,29].
2. Seed sterilization and plant preparation [30].
Critical: For a complete Pi imaging session, prepare B. distachyon plants expressing the Pi sensor and four controls simultaneously in the same manner.
a. Place up to 50 B. distachyon seeds of the same genotype in 14 mL culture tubes.
b. Add 10 mL of 20% bleach (v/v) and shake vigorously by hand for 7 min.
c. (Under the laminar hood) Decant the bleach and rinse the seeds with sterile deionized water five times.
d. Place a sterile germination paper disc in a Petri dish, wet it with sterile deionized water, and place seeds on it (seed embryo facing paper).
e. Wrap the Petri dish with Parafilm and aluminum foil and store the seeds in the dark at 4 °C for 1 week.
f. After 1 week, move the Petri dish to room temperature for 2 days to stimulate root growth, but leave the seeds in the dark (Petri dish covered with foil is suitable).
g. Take the plates out of the dark (remove foil) and place them under light (12:12 h light/dark) for 3–5 days. The light intensity in our chamber was 150 μmol/m2·s. Then, open the plates to harden the seedlings.
The B. distachyon seedlings, with a shoot length of approximately 7 cm and a root length of about 5 cm, are now ready for planting and inoculation.
3. AM fungal inoculation setup
There are many ways to grow and inoculate B. distachyon plants; here, two types of growth systems are used for the biosensor-based analysis of intracellular Pi in mycorrhizal roots:
Preparation of growth system 1: plants in cylinder cones or pots
Fill 20 cm long plastic cones or 9 cm diameter pots with growth substrates. For the cone, fill to 5 cm below the top of the cones. For the 9 cm diameter pots, fill the substrate roughly up to half the height of the pot.
Note: Choose a suitable growth substrate that allows easy access to clean roots and nutrient management. Substrates or a mixture of substrates such as sand, gravel, and/or turface with particle sizes varying from 0.05 to 0.7 mm are recommended. Avoid vermiculite and perlite as these substrates stick to the surface of the roots, which disturbs imaging. We use gravel and yellow sand in an approximately 1:1 ratio.
a. Place the AM fungal spores or the mixed propagule inoculum mixture onto the substrate and then fill the cones or the pots with the same substrate.
b. Transplant the B. distachyon seedlings into the substrate. Cover the cones/pots with a plastic dome cover for humidity control (Figure 1).
Figure 1. Demonstration of the growth system 1
c. Place the plants in a growth chamber or a greenhouse under a 12:12 h, 24:22 °C light/dark cycle. The light intensity in our chamber was 150 μmol/m2·s. Grow plants for four weeks watering with deionized water as needed. Fertilize once per week with 10 mL of 1/4 strength modified Hoagland solution containing 20 μM potassium phosphate (or as needed depending on the growth substrate used).
Note: High Pi levels in the fertilizer may inhibit fungal colonization of the host roots.
d. One day before imaging, provide plants with 10 mL of 1/4 strength modified Hoagland solution supplemented with 2 μM potassium phosphate (generating a lower Pi environment).
Preparation of growth system 2: plants inoculated between cellulose membranes
a. Sandwich a seedling and some fungal spores between two pieces of MF Millipore membrane filter discs. Position the spores around the root (Figure 2A).
Figure 2. Demonstration of the growth system 2. A. A Gigaspora gigantea spore placed next to the seedling during the system setup. B. Agar blocks placed onto the fungal hyphae to start localized treatment.
b. Place the sandwich with plants and spores vertically in the earlier discussed sand/gravel mix in a 9 cm diameter pot.
c. Set the plants in the same growth condition as described in step c in the section “Preparation of growth system 1”.
B. Preparation of B. distachyon roots for Pi imaging
Note: This procedure is tailored for Pi imaging using an upright microscope. If an inverted microscope is used, adjustments will be needed.
Growth system 1
1. Carefully remove the plants from the cones and place the root in a large Petri dish (14 cm diameter).
2. Rinse the roots gently with running water to remove substrates.
3. Immerse the roots in the container with 1/4× modified Hoagland solution with 2 μM potassium phosphate (the same Pi concentration as the final fertilizer treatment as reported in step d in the section “Preparation of growth system 1”).
Critical: To minimize the intracellular Pi disturbance due to manipulation, ensure the Pi concentration in the container matches the solution last used to maintain the plants unless another treatment is required.
4. Use a fluorescence stereomicroscope to locate a colonized root.
Note: While checking the root under the microscope, use tweezers to isolate a single root branch and gently slide it onto the glass slide.
5. Transfer the root region to the center of a microscope slide (ideally 75 mm × 25 mm). Cover it with a coverslip (ideally 20 mm × 20 mm) to maintain stability and flatness during imaging (Figure 3). If necessary, add a few drops of the 1/4 modified Hoagland solution with 2 μM potassium phosphate on the side of the coverslip.
Note: If AM fungal colonization is not required, omit steps B4 and B5, directly place a root region of interest onto the microscope slide, and cover it with a coverslip as described.
Figure 3. Arrangement of the root on the glass slide for confocal imaging
6. Gather the rest of the attached root and shoot around the edge of the coverslip (some may hang outside the slide).
7. Frequently spray the root and shoot with 1/4 modified Hoagland solution with 2 μM potassium phosphate to maintain moisture.
Growth system 2
This growth system allows a local application of Pi or chemical treatment directly to mycorrhizal roots or their connected extraradical hyphae of the fungi. The treatment is applied via solid agar blocks. Below is an example of using the system to provide a local Pi treatment of 200 μM Pi.
1. Prepare a 1/4× modified Hoagland solution containing 200 μM Pi and 1% (w/v) agar immediately before the experiment. Agar is dissolved by heating in a microwave.
Note: Sterile Hoagland solution is recommended to avoid contamination.
2. Pipette the solution into a Petri dish in 10 μL droplets and allow them to solidify.
3. Gently remove the MF Millipore membrane with the plant from the pots and place it in a ~20 cm diameter flat-bottomed container.
4. Carefully remove one MF Millipore membrane and expose the roots and fungus. Gently remove the substrates on the membrane.
Critical: Some roots grow into or through the MF Millipore membrane. Make sure the removal process does not break the roots.
5. Locate the colonized root regions and their connecting fungal hyphae. If using non-colonized roots, locate and mark the root regions of interest. Mark the site for applying the agar blocks and take photographs with distance scales for reference.
6. Place the solidified agar blocks over the roots or the extraradical hyphae at the marked sites (Figure 2B). If incubation is needed, place the membrane with the plant inside a closed container, ensuring that the shoot remains outside of the container. For example, use a 14 cm Petri dish with one side removed. The shoot can be left outside. Periodically spray the membrane with 1/4 modified solution without Pi to maintain a moist environment. The system can be left in the dark or light as needed. When applying a localized treatment to the extraradical hyphae, it may be useful to have a Benomyl-treated control. Benomyl is a fungicide, and fungal activity can be abolished by applying 1 mL of 100 μg/μL Benomyl to the mycorrhizal root system one day before the experiment (as used in [22]). This Benomyl-treated control enables an assessment of diffusion along the hyphae vs. active transport by the fungus.
7. Remove the agar blocks after 24 h of incubation. If imaging is required after the treatment, transfer the membrane with the plant onto a glass slide (ideally 75 mm × 25 mm), ensuring that the marked region is at the center of the slide.
8. Place a coverslip (ideally 20 mm × 20 mm) over the region of interest on the membrane. If necessary, add a few drops of the 1/4 modified Hoagland solution without phosphate on the side of the coverslip. Frequently spray the membrane with Hoagland solution without phosphate to maintain moisture throughout the procedure.
C. Confocal imaging of roots
Critical: See General note 1 to determine how to set the confocal settings.
1. Once confocal settings are finalized, begin imaging roots expressing either the Pi or control biosensor. Each image set should include CC (CFP excitation—CFP emission), CY (CFP excitation—YFP emission), and YY (YFP excitation—YFP emission) images. For roots generated using growth system 2, position the glass slides with the membrane directly on the microscope stage. When imaging a colonized root, ensure that the images capture the targeted arbuscules and that the arbuscules are distinct.
Note: Aim for a minimum of six biological replicates (individual roots) in each treatment group to ensure robust statistical analysis. Ideally, obtain at least five clear images showcasing at least five arbuscules in the desired colonized region for each root.
Critical: To ensure integration with our FLIPPi image analysis macro in ImageJ, maintain specific image file names when saving image files.
The suggested file naming system is pPT7_cytFLIPPi_053019_HP_0.5h_102
(sensor name) (date) (group) (plant & image number)
You can adjust the name and order of components, such as the line name, date, and treatment group, as needed. However, the plant and image number “_102” format must be retained. In this format, “1” signifies the first root, while “2” indicates it is the second image of that root. For instance, for the 10th image of root 6, the plant and image number would be “_610.” This format provides a unique identifier, allowing the FLIPPi macro to distinguish individual images from each plant.
2. After imaging all samples with the Pi and control biosensors, proceed to image the roots expressing eCFP and cpVenus. Each set of images should contain all CC, CY, and YY images.
Note: At least three biological replicates are recommended for eCFP and cpVenus, capturing a minimum of five distinct images per replicate.
D. Image processing and sensitized FRET analysis
Sensitized FRET ratios eliminate non-FRET emissions such as CFP bleed-through and YFP (or cpVenus) cross-excitation. Each image requires standard processing. The sensitized FRET analysis can be conducted for each image set (CC, CY, and YY). For efficiency, utilize the FLIPPi macro, which is designed primarily for images from Leica confocal microscopes. For other confocal types, adjust file names to fit the macro as suggested in step C1. Once the folders are directed to the FLIPPi macro, it automatically processes images and calculates CC, CY, and YY values after selecting the regions of interest (ROIs) (Figure 4). Usually, up to five ROIs can be selected per image.
Note: The macro is compatible with both MacOS and Microsoft operating systems.
Figure 4. Screenshot showcasing a sample folder with FRET images, adhering to the suggested file naming system
1. Open Fiji, find Plugins menu, go to Macros → Startup Macro. A window (Figure 5) will show up.
Figure 5. Screenshot showing the window following the selection of the Startup Macro option
2. Remove the green script as they are the default description from Fiji. Insert the command lines from Method S1 in [22]. Click Run. A subsequent window (Figure 6) will appear.
Figure 6. Screenshot showing the window following clicking Run
3. Keep the “File Name [Blank]” at its default setting. Choose one of the four functions: Series Analyzer, Reverse Series Analyzer, ROI Analyzer, or Reverse ROI Analyzer.
The functionalities of these options are detailed below:
• Series Analyzer: This function aids in background removal. It generates a mask image that captures only the root, excluding the background. This is based on the root’s profile on CC, as these images clearly depict the root profile. Subsequently, this mask is applied to the CC, CY, and YY to remove their backgrounds. The processed images are exported into the same folder as the original images.
• Reverse Series Analyzer: This operates similarly to the Series Analyzer, but with one distinction. It employs YY to create mask images instead of CC. This is specifically useful for background removal in a control set of images from cpVenus because the fluorescence intensity of CC is not high enough for mask creation.
• ROI Analyzer: After opening the processed CC created by the Series Analyzer, this function prompts users to select their regions of interest (ROIs). It then calculates the mean grey values of these ROIs from the CC, CY, and YY images. The resultant data is in tabular form, ready to be transferred into Excel.
• Reverse ROI Analyzer: Serving a similar purpose to the ROI Analyzer, this function is specifically tailored for images from the cpVenus control. It begins by opening the processed YY in lieu of the CC.
4. Begin with unprocessed images exported from the imaging software. Select Series Analyzer and click OK. This action will prompt a file directory window to appear (Figure 7).
Figure 7. Screenshot showing the file directory window after step D4
Critical: Ensure that the images exported are in TIFF format. Each set of images, originating from a single ROI, should be separated into three grayscale images: CC, CY, and YY, which are categorized based on the emission collection channels. It is important that you do not use RGB format during image processing. As specified in the macro scripts (refer to Method S1 in [22]), CC is linked to channel 00 (file names conclude with “_ch00” by Leica LAS X), CY is associated with channel 01 (indicated by “_ch01”), and YY corresponds to channel 03 (“_ch03”). The macro scripts can be tailored to various naming systems.
Critical: see General note 2.
5. Select the folder that contains your original images and click Open. All images within this folder will be automatically opened, processed, and saved back to the same location.
6. Once processing is complete, navigate to the chosen folder. Transfer the processed images to the designated “processed image” folder. You can identify the processed images by their file name endings: “-m.tif” for mask images and “-result.tif” for fully processed images. Sort the files by their date modified; all processed images are placed together (Figure 8).
Figure 8. Screenshot showing the file directory window for step D6. Processed images end their file names with “-result.tif” or “-m.tif.”
7. Run the macro again, select ROI Analyzer, and click OK.
8. Similar to the Series Analyzer, the ROI Analyzer will sequentially open each image based on the file order in the folder. An image window will appear, accompanied by a prompt asking, “How many ROI’s would you like to use?” (Figure 9).
Figure 9. Screenshot showing the prompted windows for step D8
9. Depending on the number of visualized colonized cells, typically 1–5 ROIs are chosen for each image. If an image should be bypassed, enter “0” and click OK.
Note: Adjust the brightness and contrast of the images by navigating to Image → Adjust → Brightness/Contrast… This will enhance the visibility of the ROI without altering the grayscale value of each pixel.
10. Select the Oval Tool from the taskbar and begin designating oval ROIs. Each ROI should be positioned entirely within one cell. Choose an area where the signal is bright but not saturated. Once you have chosen the ROI, click OK in the pop-up Waiting window (Figure 10).
Critical: To generate accurate data, it is essential to maintain a consistent ROI size.
A Result window will appear, displaying the CC, CY, and YY values. Gather these results once the ROIs from all images in the folder have been selected.
Figure 10. Screenshot showing the prompted windows for step D10
11. A subsequent Waiting window will emerge for the next ROI. Simply reposition the circle to another colonized cell in focus and continue the process. The macro will proceed to the next image once the designated number of ROIs has been selected from the current image.
Critical: Ensure that the ROI is selected prior to clicking OK. Failing to do so may cause the program to freeze, requiring a restart from the beginning.
12. Once the ROI Analyzer has completed analyzing all processed images in the folder (Figure 11), copy the results and paste them into the Excel template (refer to Table S4 in [22]). Close the Results window.
Figure 11. Screenshot showing the prompted windows for step D13
13. Continue the analysis for all the roots associated with FLIPPi, control sensors, and the eCFP control.
14. For roots associated with cpVenus, utilize the Reverse Series Analyzer and Reverse ROI Analyzer. The steps for using the Reverse Series Analyzer and Reverse ROI Analyzer mirror those of the Series Analyzer and ROI Analyzer, respectively.
15. Arrange the data within the Excel sheet template (refer to Table S4 in [22]). This template offers a structure where different treatment groups have individual tabs. Inside each tab, data are organized according to the sensor type. The columns labeled “Plant, CC, CY, and YY” are directly derived from the FIJI macro.
16. For CFP controls, transfer the columns titled “Plant,” “CC,” and “CY” to Excel. For cpVenus controls, transfer the columns “Plant,” “CY,” and “YY” to Excel.
Data analysis
Sensitized FRET represents a corrected CY value, where emissions not derived from FRET, such as CFP bleed-through and YFP (or cpVenus) cross-excitation (in both channels), are eliminated [20,21]. The equation for calculating sensitized FRET is:
Sensitized FRET = CY - CC*b - YY(c - a*b)
The coefficients a, b, and c are derived from the CFP and cpVenus controls, using ROI data and linear regression. A trendline from the linear regression illustrates the fit of each data point to the linear model (Figure 12). Specifically:
a represents the slope of CC against YY from cpVenus.
b is the slope of CY against CC from CFP.
c is the slope of CY against YY from cpVenus.
Figure 12. Example graph showing the trendline in Excel
These coefficients can be determined using Excel’s regression functions on a graph or the LINEST function [23,24,31].
Typically, the coefficient a is close to zero, allowing the calculation to be simplified to:
Sensitized FRET = CY - CC*b - YY*c
Before utilizing the simplified equation, it is important to verify the value of a. Subsequently, FRET ratios can be derived by dividing the sensitized FRET by CC. Use linear regression graphs to evaluate the linearity of the sensitized FRET. A closer alignment of data to the linear model signifies greater accuracy of the sensitized FRET result.
For a reliable and in-depth statistical assessment, it is advisable to have at least six biological replicates.
Validation of protocol
This protocol has been used and validated in the following research article:
• Zhang et al. [22]. A genetically encoded biosensor reveals spatiotemporal variation in cellular phosphate content in Brachypodium distachyon mycorrhizal roots. New Phytol 234: 1817–1831. https://doi.org/10.1111/nph.18081
General notes and troubleshooting
General notes
1. Prior to imaging, optimizing the confocal settings (such as laser, objective, filter, and scan mode) is critical for generating reliable and consistent FRET data. The goal is to obtain a high range of fluorescence intensities, but no pixels should be saturated. To achieve this goal, a few transgenic plants expressing the Pi biosensor and its three controls need to be included in the process. After the settings are confirmed, they should be kept the same across all control lines for the same Pi biosensor. If imaging tissue types and biosensor lines are changed, the optimizing process needs to be redone. The following criteria need to be considered:
• For the sensitized FRET analysis, each set of the root image from the Pi and control biosensor needs to include three images: CFP excitation—CFP emission (CC hereafter), CFP excitation—YFP emission (CY hereafter), and YFP excitation—YFP emission (YY hereafter). When capturing CC and CY, maintain the exact same confocal settings, such as the excitation wavelength and intensity, exposure time, scan speed, gain, and pinhole (if any is applicable). Ensure the fluorescent intensities of YY remain roughly equivalent to those in CC and CY.
• Because CC and CY emission collection channels are very close on the wavelength, try to narrow the two channels as much as possible to avoid bleed-through but also keep a reasonable fluorescent intensity.
• Images of non-fluorescent wildtype B. distachyon need to be included for background subtraction purposes.
• Laser power. Some lasers need warm-up time; thereby, the laser power may increase during imaging. Consider warm-up time before starting imaging.
For our experiments, the objective used for plants in growth system 1 expressing the BdPT7-cytosolic, ZmUbi-cytosolic, and BdPT7-plastidic sensors was a 63× water lens with a numerical aperture of 1.20, whereas a 20× water lens with a numerical aperture of 0.70 was used for plants growing in growth system 2. The 458 nm laser intensity (used for CC and CY) was 40% for BdPT7-cytosolic sensors, 90% for ZmUbi-cytosolic sensors, and 30% for BdPT7-plastidic sensors. The 514 nm laser intensity (used for the YY) was 10% for BdPT7-cytosolic sensors, 16% for ZmUbi-cytosolic sensors, and 7% for BdPT7-plastidic sensors. The emission ranges collected by the HyD detector for the CC channel were between 475 and 490 nm, while for the CY and YY channels, the emission range was between 546 and 562 nm. The HyD detector operated in photon counting mode for all conditions. Additional parameters included a line average of 3, a pinhole setting of 1 Airy Unit, a scan speed of 400 Hz, and a sequential scan mode set between frames. These settings were used consistently across the different sensor conditions.
2. For a more efficient workflow, it is recommended to structure the organization of the files as follows, which aligns with the macro's parameters:
Folder: xxxx [experiment name] original images
(This main folder should house the initial images directly exported from the imaging software.)
Within this, separate folders for original images based on their genotypes. The folder names can differ, but they should be grouped based on genotype:
Sub-folder 1: FLIPPi
Sub-folder 2: Null
Sub-folder 3: CFP
Sub-folder 4: cpVenus
For better organization, include folders to house processed images by the Series Analyzer/Reverse Series Analyzer:
Sub-folder 5: FLIPPi-processed
Sub-folder 6: Null-processed
Sub-folder 7: CFP-processed
Sub-folder 8: cpVenus-processed
Troubleshooting
Problem 1: The CY signals are very low.
Possible cause: The laser power is too low, or minimal signals are being captured in the CY channel.
Solution: Boost the laser power, enhance the digital signal collection gain, or expand the emission collection wavelength range.
Problem 2: Inconsistent FRET observed among replicates of the same genotype.
Possible cause: The confocal setting is not optimized.
Solution: Reoptimize the confocal settings; ensure that the fluorescence intensities for both the Pi sensor and controls are robust yet not saturated.
Problem 3: The Fiji FLIPPi macro fails to read the image files correctly.
Possible cause: The file names are not recognized by the macro.
Solution: Ensure the file naming aligns with the recommendations provided in the procedure. Thoroughly check for any typographical errors, spaces, or underscores in the file names.
Acknowledgments
This protocol was used in Zhang et al. [22]. Funding for this research was provided by the US Department of Energy, Office of Science, Office of Biological and Environmental Research (grant no. DE-SC0014037). SZ's work was partially supported by NIFA postdoctoral fellowship-converted standard grant (Award Number # 2021-67034-39677). The confocal microscope utilized in the BTI Plant Cell Imaging Center was acquired through a US NSF Instrumentation Grant, DBI-0618969. We thank the members of Dr. Wayne K. Versaw’s lab at Texas A&M University for insightful discussions and the BTI Computational Biology Center for guidance on using the Fiji image processor.
Competing interests
The authors declare no competing financial interests.
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Article Information
Publication history
Received: Sep 4, 2024
Accepted: Nov 10, 2024
Available online: Dec 3, 2024
Published: Jan 20, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Biochemistry > Protein > Fluorescence
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The authors used this protocol in eLIFE May 2024
Abstract
Mouse embryonic fibroblasts (MEFs) derived from genetically modified mice are a valuable resource for studying gene function and regulation. The MEF system can also be combined with rescue studies to characterize the function of mutant genes/proteins, such as disease-causing variants. However, primary MEFs undergo senescence soon after isolation and passaging, making long-term genetic manipulations difficult. Previously described methods for MEF immortalization are often inconsistent or alter the physiological properties of the cells. Here, we describe an optimized method that overcomes these limitations. By using electroporation to deliver CRISPR constructs that target the Tp53 gene, the method reliably generates immortalized MEFs (iMEFs) within three weeks. Importantly, iMEFs closely resemble the parent cell populations, and individual iMEFs can be cloned and expanded for subsequent genetic manipulation and characterization. We envision that this protocol can be adopted broadly to immortalize other mouse primary cell types.
Key features
• CRISPR-based knockout of the Tp53 gene enables efficient immortalization of mouse embryonic fibroblasts (MEFs) in under three weeks.
• Immortalization requires a Neon electroporator or a comparable system to transfect cells with the Tp53 CRISPR constructs.
Keywords: Mouse embryonic fibroblasts (MEFs) Immortalized mouse embryonic fibroblasts (iMEFs) MEF isolation MEF immortalization Tp53 knockout CRISPR Electroporation
Background
The ability to culture cells long-term in vitro has been a cornerstone of modern cell biological research. Except for stem and cancer cells, most primary cells undergo senescence in culture, making immortalization an essential step during the establishment of permanent cell lines from native tissues. Earlier methods of cell immortalization involved overexpression of oncogenes to transform cells [1–4]. Though efficient, these methods frequently result in the acquisition of cancer-like phenotypes, including the loss of contact inhibition and anchorage-dependent growth, as well as alterations in growth factor requirement, metabolism, and other signaling activities [5–7].
Overexpression of telomerase (TERT, or telomerase reverse transcriptase) is another common method to transform mammalian cells [8]. This method is highly effective in immortalizing human cells and has a lower tendency to induce cancer-like phenotypes [9]. However, few examples exist in the literature demonstrating that TERT overexpression alone can immortalize mouse cells [10], suggesting that this method is generally less effective toward mouse cells. In our own experience, we have not been able to successfully immortalize mouse embryonic fibroblasts via overexpression of telomerase.
3T3 cells are widely used mouse embryonic fibroblast (MEF) lines spontaneously immortalized through serial passaging. The “3T3” method, originally described by Howard Green and colleagues, involves seeding primary MEFs at 3 × 105 cells per 50-mm dish and transferring every 3 days at the same density until rapidly dividing, immortalized cells emerge [11]. While this immortalization process is considered gentle and does not cause neoplastic transformation [11], it is inefficient and time-consuming. Moreover, because the method relies on mutation(s) that spontaneously arise during the prolonged cell passaging phase, it is difficult to compare cell lines derived from different immortalization experiments.
Notably, Harvey and Levine [12] reported that MEF lines immortalized via the 3T3 method frequently carry loss-of-function mutations in the tumor suppressor gene Tp53, suggesting that this is a key immortalizing event. Consistent with this observation, we found that CRISPR-mediated ablation of the Tp53 gene robustly immortalizes primary MEFs. We have used the protocol described here to reliably generate immortalized MEF lines (iMEFs) in under three weeks and from as few as 50,000 primary cells. iMEF lines have also been successfully established from wildtype (WT) and genetically modified embryos of different genetic backgrounds (C57BL/6 and mixed C57BL/6; 129J). Lastly, iMEFs generated through Tp53 deletion can be subcloned and genetically manipulated further in downstream applications, such as gene rescue experiments.
Materials and reagents
Biological materials
1. Pregnant mice carrying embryonic day 12.5 (E12.5) embryos
Reagents
1. Phosphate buffered saline (PBS) without Ca2+ and Mg2+ (Cytiva, catalog number: SH30256.01)
2. Hanks’ balanced salt solution (HBSS) without Ca2+ and Mg2+ (Gibco, catalog number: 14170112)
3. Trypsin from bovine pancreas (Worthington Biochemical, catalog number: LS003702) for MEF preparation
4. RQ1 RNase-free DNase (1 u/μL) (Promega, catalog number: M6101)
5. Non-essential amino acids solution in minimal essential medium (MEM NEAA) (100×, Gibco, catalog number: 11140050)
6. Sodium pyruvate (100 mM) (Gibco, catalog number: 11360070)
7. Dulbecco’s modified Eagle’s medium (DMEM), high glucose (4.5 g/L), 1× (Gibco, catalog number: 11960-044)
8. Fetal bovine serum (FBS), United States qualified grade (Gibco, catalog number: 26-140-079; this product has been discontinued, but other similar grade FBS can be used)
9. Penicillin-streptomycin (100×) (Gibco, catalog number: 15-140-122)
10. L-glutamine (200 mM) (Gibco, catalog number: 25030081)
11. Dimethyl sulfoxide (DMSO), sterile, tissue culture grade (Millipore Sigma, catalog number: D2650-100ML)
12. 0.25% trypsin-EDTA with phenol red (Gibco, catalog number: 25200072), for cell harvest and passaging after the initial MEF preparation step
13. Px461-Cas9n-Trp53-sgRNA-alpha plasmid (Addgene, plasmid number: 88846; generated and deposited by the Massagué lab) [13]
14. Px461-Cas9n-Trp53-sgRNA-beta plasmid (Addgene, plasmid number: 88847; generated and deposited by the Massague lab) [13]
15. pCAG-GFP plasmid (Addgene, plasmid number: 11150; generated and deposited by the Cepko lab) [14]
Solutions
1. 0.1% trypsin for MEF preparation (see Recipes)
2. Complete cell culture media (culture media) (see Recipes)
3. Cell freezing media (see Recipes)
Recipes
1. 0.1% trypsin for MEF preparation
Reagent Final concentration Amount
Trypsin from bovine pancreas 0.1% 40 mg
HBSS 1× 40 mL
Filter sterilize with a 0.22 μm syringe filter. Aliquot, snap freeze in liquid nitrogen, and store at -80 °C.
2. Complete cell culture media (culture media)
Reagent Final concentration Amount
DMEM 1× 500 mL
FBS 10% 50 mL
Penicillin-streptomycin (100×) 1× 5.5 mL
L-glutamine (200mM) 2mM 5.5 mL
Prewarm at 37 °C before use. For culturing freshly isolated MEFs (steps A2k–o), supplement culture media with sodium pyruvate (1mM final) and MEM NEAA (1× final). For culturing cells immediately after electroporation (steps B3–19), use culture media without penicillin-streptomycin.
3. Cell freezing media
Reagent Final concentration Amount
Culture media 90% 45 mL
DMSO 10% 5 mL
Filter sterilize with a 0.22 μm syringe filter.
Laboratory supplies
1. Surgical scissors (Fine Science Tools, model: Surgical scissors-sharp, catalog number: 14002-12)
2. Adson forceps (Fine Science Tools, model: Adson forceps, catalog number: 11006-12)
3. Fine tip forceps (Fine Science Tools, model: Dumont #5 Inox, catalog number: 11251-20)
4. 60 mm tissue culture (TC)-treated cell culture dishes (Corning, Falcon, catalog number: 353002)
5. 100 mm TC-treated cell culture dishes (Corning, Falcon, catalog number: 353003)
6. 6-well clear TC-treated multiple well plates (Corning, Costar, catalog number: 3516)
7. 24-well clear TC-treated multiple well plates (Corning, Costar, catalog number: 3524)
8. 50 mL conical centrifuge tubes (Thermo Fisher Scientific, Nunc, catalog number: 339653)
9. Pipettes (Eppendorf, Research Plus, P-1000, P-200, P-20, catalog number: 3123000918)
10. Aerosol barrier pipette tips [Fisher Scientific, SureOne, catalog number: 02-707-442 (0.1–10 μL), 02-707-432 (2–20 μL), 02-707-430 (20–200 μL), 02-707-404 (100–1,000 μL)]
11. Metal tube rack for 1.5 mL tubes (Stratagene, catalog number: 41018; the item has been discontinued, but any similar rack will work)
12. 5 mL serological pipettes (Thermo Fisher Scientific, Nunc, catalog number: 170355N)
13. Pipette controller (Drummond, Pipet-Aid, catalog number: 4-000-100)
14. 1.5 mL microcentrifuge tubes (Denville Scientific, PosiClick, catalog number: C2170)
15. Neon transfection system 10 μL kit (Thermo Fisher Scientific, Neon, catalog number: MPK1096); includes Neon tips, Neon tubes, buffer R, and buffer E
16. TransIT-LT1 transfection reagent (Mirus Bio, catalog number: MIR 2304)
17. Cell strainers (Corning, Falcon, catalog number: 352340)
18. 30 mL syringes (BD, catalog number: 302832)
19. 0.22 μm syringe filters (Thermo Fisher Scientific, Fisherbrand, catalog number: 09-719C)
20. Cryogenic vials (Corning, catalog number: 430488)
Equipment
1. Dissecting microscope (Leica, model: S7E, catalog number: S7E-PS)
2. Biological safety cabinet (Thermo Fisher Scientific, model: 1300 series class II, catalog number: 1323TS)
3. CO2 incubator (Thermo Fisher Scientific, model: Heracell 150i, catalog number: 51026281), set at 37 °C with 5% CO2 and 90%–95% humidity
4. Benchtop centrifuge (Thermo Fisher Scientific, model: Sorvall ST 8, catalog number: 75007200)
5. 8 × 50 swinging bucket rotor (Thermo Fisher Scientific, model: TX-100S, catalog number: 75005704)
6. Microcentrifuge (Thermo Fisher Scientific, model: Sorvall Legend Micro 21, catalog number: 75002436)
7. 24 × 1.5/2.0 mL rotor with ClickSeal lid (Thermo Fisher Scientific, catalog number: 75003424)
8. Ultra-low (-86 °C) freezer (Thermo Fisher Scientific, model: Revco UxF, catalog number: UXF60086A; this model has been discontinued)
9. Liquid nitrogen storage unit (Thermo Fisher Scientific, model: Cryoplus 3, catalog number: 7404)
10. Cell counter (Corning, catalog number: 6749)
11. Electroporation device and Neon pipette (Thermo Fisher Scientific, model: Neon, catalog number: MPK5000. This instrument has been discontinued; other similar electroporation devices can be used, but optimization will be required.)
12. Inverted fluorescence microscope (Thermo Fisher Scientific, model: Evos FL, catalog number: AMF4300)
13. Mr. Frosty freezing container (Thermo Fisher Scientific, catalog number: 5100-0001)
Procedure
A. Preparation of primary mouse embryonic fibroblasts (MEFs)
Performing the embryo dissection in a laminar flow hood is ideal. However, if a laminar flow hood is not available, the procedure can be performed on a standard lab bench. All solution stocks (e.g., HBSS) should be kept sterile and transferred to the dishes used for dissection in a biosafety cabinet. General procedures on mouse husbandry can be found in [15].
1. Dissection of E12.5 embryos from timed mating
a. Set up timed mating and start checking the females for vaginal plugs the following morning.
Critical: Plugged females should be separated from the male and housed until 12.5 days post-coitum.
b. Euthanize the dam at 12.5 days post-coitum via CO2 inhalation followed by cervical dislocation to ensure death.
c. Spray the dam with 70% ethanol to sterilize the abdominal surface.
d. Dissect out the uterine horns and rinse in PBS in 100 mm dishes (Figure 1A–1B).
Critical: When dissecting the uterine horns, ensure that the intestines are not nicked and the uterine horns do not touch the fur to prevent contamination.
e. Separate the embryos from the uterus and place each embryo in a 60 mm dish containing HBSS (Figure 1C–1D).
f. In the same 60 mm dish and under a dissecting microscope, remove the placenta and embryonic sac (Figure 1D–1E).
Note 1: The embryonic sacs can be collected and used to genotype the embryos. One-half to one-quarter of each sac is sufficient.
Note 2: The sac should be gripped firmly with fine-tip forceps and rinsed briefly under a slow stream of running deionized water before transferring to a collection tube. This helps to prevent cross-contamination from maternal tissues.
g. Remove the brain, eyes, and internal organs of the embryo as much as possible using fine-tip forceps and discard them (Figure 1E–1F).
h. Transfer the dissected embryo to a 6-well plate filled with HBSS and keep the plate on ice until all embryos have been dissected. Place one dissected embryo in each well of the 6-well plate.
Critical: The plate(s) should be kept on ice to ensure the viability of embryonic tissues. Proceed to step A2 as soon as possible.
Note: The carcass of the dam and other maternal and embryonic tissues should be disposed of through approved biohazard protocols.
Figure 1. Key steps of embryo dissection. A. Top panel: the approximate position of the uterus in the dam. Bottom panel: dissected dam showing the uterus with embryos. B. A dissected uterine horn with embryos in it. C. An embryo still wrapped in the transected uterine wall. D. An embryo still encased in the embryonic sacs. * denotes the placenta. E. An embryo after the removal of the embryonic sacs and placenta. F. An embryo after the removal of the brain, eyes, and internal organs.
2. Preparation of MEFs
This and all subsequent steps should be performed in a biosafety cabinet under sterile conditions.
Before starting:
Prewarm the metal rack at 37 °C in the TC incubator.
Thaw 0.1% trypsin (from bovine pancreas) at 37 °C and keep it on ice once fully thawed.
Label 1.5 mL microcentrifuge tubes for embryo homogenization. Label as many tubes as there are embryos.
a. Add 25 μL of RQ1 DNase into the first labeled 1.5 mL tube. Add 0.5 mL of 0.1% trypsin to the same tube and mix by pipetting 2–3 times with a P1000 pipette.
b. Transfer the first embryo from the 6-well with HBSS into the tube (containing trypsin and DNase I) using a P1000 pipette while applying gentle suction.
c. Homogenize the embryo by pipetting up and down 12–15 times using a P1000 pipette until the embryo is broken into small chunks (Figure 2A).
d. Store the tube with the homogenized embryo on ice and proceed to homogenize the next embryo (repeating steps A2a–A2c), until all embryos are homogenized.
Critical:
Prepare each tube of RQ1 DNase and trypsin after processing the previous embryo. This ensures optimal activity of the DNase.
It is very important to not over-homogenize the embryo as this can cause excessive cell lysis. When the homogenate starts to feel increasingly viscous, it is over-homogenized.
e. Transfer the tubes with the homogenized embryos to the prewarmed metal rack and incubate at 37 °C for exactly 8 min.
Note: The homogenized tissue chunks tend to settle down in the tube. Invert the tube every ~3 min to mix the contents of the tube.
f. While the incubation is ongoing, add 5 mL of cold culture media to a 50 mL conical tube in preparation for the next step. Prepare as many tubes as there are embryos.
Note: The media should be cold to facilitate rapid neutralization of the trypsin.
g. After the 8-min incubation at 37 °C, transfer 1 mL of the cold culture media from the 50 mL tube (step A2f) to the 1.5 mL microcentrifuge tube containing the homogenate and trypsin (Figure 2B) to neutralize the trypsin.
h. Pipette gently 2–3 times with a P1000 pipette and transfer the contents (Figure 2C) into the 50 mL tube with the remaining 4 mL cold culture media (from step A2f).
i. Use a 5 mL serological pipette to pipette the mixture up and down 8–10 times very gently.
j. Centrifuge the neutralized homogenates at 500× g for 10 min at room temperature in a centrifuge equipped with a swinging bucket rotor.
k. While centrifugation is ongoing, add 10 mL of prewarmed culture media supplemented with 100 μL of sodium pyruvate (1 mM final concentration) and 100 μL of MEM NEAA (1× final concentration) to a 100 mm TC-treated cell culture dish. Prepare as many dishes as there are embryos. Keep the dishes in the TC incubator.
l. Once the centrifugation is done, carefully remove the supernatant by aspiration.
Critical: Be careful to avoid disturbing the pellet (Figure 2D) or getting the aspirator tip too close to the pellet, as the pellet can easily get aspirated away.
m. Gently resuspend each pellet in 1 mL of culture media supplemented with sodium pyruvate and MEM NEAA (from the 100 mm dish prepared in step A2k) using a P1000 pipette and transfer the cell suspension back to the same prewarmed 100 mm dish.
n. Shake the plate gently to ensure uniform distribution of the cells.
o. Incubate the cells in the TC incubator (37 °C, 5% CO2, 90%–95% humidity) without disturbance until the next day.
p. The next morning, observe the cells to assess cell health (Figure 2E–2F).
Note 1: The MEFs should be approximately 80% confluent one day after isolation and reach full confluency within 2–3 days. Some cell death is expected, but most cells should survive and adhere to the culture dish by the morning after isolation (Figure 2E).
Note 2: It is normal for the MEF cultures to contain clumps of tissue that are not fully dissociated (Figure 2F). Leave the clumps as they are.
q. Once the cells are confluent, wash once with sterile PBS and trypsinize the cells in each 100 mm dish with 3.5 mL of prewarmed 0.25% trypsin-EDTA.
r. Once the cells lift off the plate (~1–2 min), neutralize the trypsin with 7 mL of culture media.
s. Filter the cell suspension (still in the trypsin/culture media mixture) through a 40 μm cell strainer and collect the flowthrough in a 50 mL tube.
Note: This step removes any remaining large cell/tissue clumps.
t. Centrifuge the cell suspension at 500× g for 5 min at room temperature in a centrifuge equipped with a swinging bucket rotor.
u. Carefully aspirate the supernatant. At this point, cells can be split and cultured further (step A2v below) or frozen (step A2w below).
v. To culture the MEFs further, resuspend the cell pellet in 1 mL of culture media and pipette 12–15 times with a P1000 pipette. Proceed with splitting into the desired number of dishes (ideally no more than a 1:5 split).
w. To freeze the MEFs, resuspend the cell pellet from each 100 mm dish in 1 mL of cell freezing media and transfer to a cryogenic vial. Smaller aliquots can be prepared if desired. Put the vials in a room-temperature Mr. Frosty freezing container and then store them in an ultra-low freezer at -80 °C overnight. For long-term storage, transfer the cryogenic vials to a liquid nitrogen freezer.
Pause point: Primary MEFs can be stored in liquid nitrogen for many years.
Caution: Precautions (eye protection, cryo-safe gloves, and lab coat) must be taken while handling liquid nitrogen.
Figure 2. Key steps of mouse embryonic fibroblast (MEF) preparation. A. Embryo homogenate before the 8-min, 37 °C incubation with trypsin. B. Embryo homogenate after the 8-min, 37 °C incubation with trypsin. C. Embryo homogenate after neutralization of trypsin with 1 mL of culture media. D. Cell pellet (indicated by the black arrow) after centrifugation of the homogenate. E–F. Representative phase contrast images of MEFs 16 h after isolation. A large clump of cells is visible in the upper left corner of (F) Scale bar: 50 μm. Magnification: 10×.
B. Immortalization of MEFs
Before starting:
Approximately 2 days before electroporation, seed a MEF stock such that cells reach 70%–90% confluency and are growing robustly on the day of electroporation. Cells can be split from an ongoing culture or thawed from a frozen stock. Cells should be under 3 passages. We routinely use passage 0 (P0) or P1 cultures. A minimum of 200,000 cells are ideal for the experiment. We typically prepare 1–2 million cells (two wells of a 6-well plate)
Prepare the plasmids required for the immortalization procedure and adjust the concentration to 1 μg/μL.
Before starting the electroporation experiment, prepare culture media without penicillin-streptomycin and add 500 μL per well to a 24-well plate; prewarm at 37 °C. Also, prewarm 0.25% trypsin-EDTA at 37 °C.
Review the Neon electroporator guide (https://assets.thermofisher.com/TFS-Assets/LSG/manuals/neon_device_man.pdf; https://www.thermofisher.com/content/dam/LifeTech/migration/en/filelibrary/cell-culture/neon-protocols.par.71910.file.dat/mouse%20embryonic%20fibroblasts%20(mef)-embryo.pdf).
1. Wash cells once with PBS (1 mL for each 6-well) and trypsinize with 0.25% trypsin-EDTA (0.3 mL for each 6-well).
2. Incubate at 37 °C until the cells lift off the plate (~1–2 min).
3. Neutralize the trypsin with culture media without penicillin-streptomycin (0.7 mL for each 6-well) and transfer the cell suspension to a 1.5 mL microcentrifuge tube or another appropriately sized tube.
4. Pipette the cells 8–10 times using a P1000 pipette to achieve a single-cell suspension.
5. Count the cells and determine the cell density (cells are still in the trypsin/culture media mixture at this point).
6. Transfer 100,000 cells into each of two 1.5 mL microcentrifuge tubes (each transfection requires 50,000 cells; the two sets of cells are prepared for Tp53 CRISPR and GFP transfections. Double the amount of cells is prepared in each set to account for pipetting errors).
7. Centrifuge the cells at 500× g for 5 min in a microcentrifuge. Carefully aspirate the trypsin/culture media mixture.
8. Resuspend the cell pellet in 0.5 mL of PBS. Centrifuge to pellet the cells (as in step B7).
Critical: This step removes the residual trypsin/culture media mixture and is critical for successful electroporation.
9. Carefully aspirate the PBS and resuspend the cell pellet in each tube in 20 μL of TransIT-LT1 transfection reagent. This is equivalent to a cell density of 5 × 106 cells/mL in the transfection reagent.
Note 1: Remove as much PBS as possible without disrupting the cell pellet, which is very small, before adding the transfection reagent.
Note 2: The TransIT-LT1 transfection reagent causes less cell death than buffer R included in the Neon transfection kit. However, we have had success with buffer R as well.
10. For the Tp53 CRIPSR and GFP control transfections, prepare a reaction mixture according to Table 1.
Note: Each electroporation reaction requires 10 μL of the reaction mixture. To account for pipetting errors, a reaction mixture with a larger volume (20 μL) is prepared. If desired, duplicates or triplicates of the electroporation procedure can be performed to safeguard against occasional suboptimal electroporation (see note under step B16).
Table 1. Recipe for the Tp53 CRISPR and GFP reaction mixtures used for electroporation
Tp53 CRISPR reaction mixture GFP reaction mixture
MEF suspension in the transfection reagent (density: 5 × 106 cells/mL) 20 μL 20 μL
pCAG-GFP (1 μg/µL) -- 0.5 μL
Px461-Cas9n-Trp53-sgRNA-alpha (1 μg/μL) 0.25 μL --
Px461-Cas9n-Trp53-sgRNA-beta (1 μg/μL) 0.25 μL --
11. Mix the cells and DNA gently by pipetting using a P20 pipette.
12. Fill the Neon tube with 3 mL of buffer E and insert into the pipette station of the Neon transfection system.
Note: The pipette station should be placed in the biosafety cabinet to ensure sterility. The pulse generator can be kept outside the biosafety cabinet.
13. Insert a Neon tip into the Neon pipette by pressing the push-button on the Neon pipette to the second stop. Firmly insert the Neon pipette into the Neon tip. Gently release the push button while still applying some downward pressure to ensure a tight fit.
14. Pipette 10 μL cell/DNA suspension from the Tp53 CRISPR reaction mixture.
Critical 1: Ensure that there are no air bubbles trapped in the tip.
Critical 2: Ensure that the cells in the master mix are in a uniform suspension. If cells have settled, mix by pipetting up and down using the Neon pipette.
15. Insert the Neon pipette into the Neon tube with buffer E.
16. Electroporate using the program below (Table 2):
Table 2. Electroporation parameters for MEF immortalization
Pulse voltage (V) Pulse width (ms) Number of pulses
1350 30 1
Note 1: A very small spark may occur during the pulse. If a large visible spark is seen, it could indicate an issue in the conductance due to a trapped air bubble.
Note 2: The electroporation parameters may need to be optimized for specific experiments, especially if embryos of a different age or genetic background are used. More information on alternate programs is available in the Neon electroporator guide (https://assets.thermofisher.com/TFS-Assets/LSG/manuals/neon_device_man.pdf).
17. Immediately transfer the cells from the Neon pipette tip into the prepared 24-well plate (prewarmed culture media without penicillin-streptomycin). Gently shake the 24-well plate to evenly distribute the cells.
18. Repeat steps B14 to B17 for the GFP control construct using the GFP reaction mixture.
Note: We routinely use each tip for up to six electroporations. However, a fresh tip can be used for each electroporation if desired.
19. Incubate the cells in a TC incubator (37 °C; 5% CO2; 90%–95% humidity).
Note: Cells can be observed for viability ~2 h after electroporation.
Critical: Plate disturbance should otherwise be kept to a minimum.
20. Evaluate the electroporation efficiency by observing the expression of GFP in the GFP control group under an inverted fluorescence microscope 16–24 h after electroporation. Some cell death (20%–60%, depending on the electroporation program used) is expected after electroporation. >30% of the healthy cells should show GFP expression (Figure 3A).
Figure 3. Representative images of mouse embryonic fibroblasts (MEFs) after electroporation and throughout the process of immortalization. A. GFP channel and phase contrast images of cells transfected with the GFP control or the Tp53 CRISPR constructs 16 h after electroporation. B. Representative phase contrast images of cells transfected with the GFP control or Tp53 CRISPR constructs. Top panels: cells imaged four days after electroporation, before differences in the cell proliferation rate are detectable between the two conditions. Bottom panels: Cells imaged 17 days after electroporation and after being split twice. Cells transfected with the GFP control construct underwent senescence (as indicated by the larger size) or died. In contrast, cells transfected with the Tp53 CRISPR constructs proliferate robustly, indicating successful immortalization. Scale bar: 50 μm. Magnification: 10×.
21. Sixteen to twenty-four hours after electroporation, change culture media to complete cell culture media (with penicillin-streptomycin).
22. Continue to culture and passage the cells when they reach confluency (Figure 3B, top panel) (we typically do a 1:15 split) until the cells in the GFP control group have died off (usually within 2–3 weeks). Cells from the Tp53 CRISPR group should continue to proliferate, which indicates successful immortalization (Figure 3B, bottom panel).
23. Freeze down the established iMEFs and/or culture for further experiments.
Note: All waste from tissue culture procedures should be disposed of through approved biohazard protocols.
Validation of protocol
We have successfully used this protocol to generate more than 15 independent iMEF lines, including those described in Griffiths et al. (Figures 3–5 and associated supplementary data figures) [16].
General notes and troubleshooting
Troubleshooting
Problem Possible cause Solution
Low cell viability after establishing MEFs The trypsin digestion is too long. Adjust the trypsinization time and neutralize the trypsin as soon as the incubation is over.
The dissected embryos are over-homogenized. Reduce the number of times that the dissected embryos are pipetted up and down. The homogenization step should be gentle.
The MEF isolation procedure takes too long. Get familiar with the procedure and have all solutions and supplies ready before starting.
Low cell viability after thawing MEFs Cells are not healthy or have already undergone senescence. If the freshly isolated MEFs from each embryo do not reach confluency within 3 days, or if excessive death is observed on the day after isolation, the culture should be discarded, and the MEF isolation should be repeated from new embryos.
Low cell viability after electroporation Poor cell health before electroporation. The cells must be healthy and proliferating robustly before electroporation.
DNA concentration is not optimal for the cells being electroporated. The concentration of DNA used for the electroporation should be optimized. Try reducing the DNA concentration.
Microbubbles in the Neon pipette tip. Ensure that the Neon tip is firmly fitted onto the Neon pipette. Press the push button down to the first stop fully before inserting the tip into the cell/DNA master mix to withdraw the sample.
Avoid introducing bubbles while mixing the cell/DNA suspension.
The electroporation program is too harsh for the cells. Try different electroporation parameters.
The electroporation buffer is not optimal for the cells Other electroporation buffers, such as Buffer R (from the Neon kit) can be used, though we have found the TransIT-LT1 transfection reagent to perform better.
Endotoxin contamination in the DNA preparation. Use an endotoxin-free purification kit for DNA isolation.
Low electroporation efficiency Electroporation parameters are too weak. Optimize by increasing pulse voltage, width, and number (see page 34 of the Neon transfection user guide:
https://assets.thermofisher.com/TFS-Assets/LSG/manuals/neon_device_man.pdf).
Cells are not proliferating robustly prior to electroporation. Ensure that the cells are within 0–3 passages and are proliferating robustly. Also, ensure that cells have not undergone senescence. (Cells in senescence usually appear larger.)
DNA preparations have salt contamination or other impurities. Use a high-quality purification kit for DNA isolation. Wash the DNA pellets with 70% ethanol during isolation to remove salt.
Acknowledgments
This study was supported by the National Institutes of Health (1R35GM144341) to H.H.H. This protocol was established to generate the iMEFs used in the following publication: Griffiths et al. [16]. We would like to thank Joan Massagué for gifting the Px461-Cas9n-Trp53-sgRNA-alpha and Px461-Cas9n-Trp53-sgRNA-beta plasmids, and Connie Cepko for gifting the pCAG-GFP plasmid. The graphical abstract was prepared using Biorender.com.
Competing interests
The authors declare no competing interests. Correspondence and requests for materials should be addressed to H.H.H. ([email protected]).
Ethical considerations
All protocols using mice have been approved by Institutional Animal Care and Use Committee (IACUC), University of California, Davis.
All protocols involving DNA technology and biohazardous materials have been approved by the Institutional Biosafety Committee (IBC) at University of California, Davis.
References
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Article Information
Publication history
Received: Jun 27, 2024
Accepted: Nov 7, 2024
Available online: Nov 28, 2024
Published: Jan 20, 2025
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© 2025 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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Stem Cell > Embryonic stem cell > Maintenance and differentiation
Cell Biology > Cell isolation and culture
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5,160 | https://bio-protocol.org/en/bpdetail?id=5160&type=0 | # Bio-Protocol Content
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An Optimized Protocol for Simultaneous Propagation of Patient-derived Organoids and Matching CAFs
JH Jenny M. Högström
TM Taru Muranen
Published: Vol 15, Iss 2, Jan 20, 2025
DOI: 10.21769/BioProtoc.5160 Views: 1770
Reviewed by: Anca Flavia Savulescuilgen Mender Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in The Journal of Biological Chemistry Aug 2023
Abstract
Recurrent hormone receptor-positive (HR+) breast cancer is a leading cause of cancer mortality in women. Recurrence and resistance to targeted therapies have been difficult to study due to the long clinical course of the disease, the complex nature of resistance, and the lack of clinically relevant model systems. Existing models are limited to a few HR+ cell lines, organoid models, and patient-derived xenograft models, all lacking components of the human tumor microenvironment. Furthermore, the low take rate and loss of estrogen receptor (ER) expression in patient-derived organoids (PDOs) has been challenging. Our protocol allows simultaneous isolation of PDOs and matching cancer-associated fibroblasts (CAFs) from primary and metastatic HR+ breast cancers. Importantly, our protocol has a higher take rate and enables long-term culturing of PDOs that retain ER expression. Our matching PDOs and CAFs will provide researchers with a new resource to study the influence of the tumor microenvironment on various aspects of cancer biology such as cell growth and drug resistance in HR+ breast cancer.
Key features
• Propagation of patient-derived organoids and matching cancer-associated fibroblasts from primary and metastatic hormone receptor (HR+) positive breast cancer.
• Optimized media for long-term culturing of HR+ organoids from primary tumors and bone metastasis.
• Co-culture model to assess the influence of the tumor stroma on breast cancer progression.
Keywords: Patient-derived organoids Cancer-associated fibroblasts Co-culture model Tumor-stroma crosstalk Hormone receptor-positive breast cancer
Graphical overview
Graphical overview of the key steps for establishing patient-derived organoid cultures and matching cancer-associated fibroblasts from hormone receptor-positive breast cancer
Background
Breast cancer is the second leading cause of cancer-related mortality in women with over 600,000 deaths yearly. The majority of newly diagnosed breast cancers are hormone receptor-positive (HR+), expressing estrogen receptor (ER) with or without progesterone receptor (PR) [1]. These patients are treated with targeted therapies, such as endocrine therapies and CDK4/6 inhibitors [2]. Although HR+ breast cancers typically respond well to these therapies, approximately 30% of the patients will relapse. Unfortunately, recurrent HR+ breast tumors are usually metastatic and incurable. Besides tumor-intrinsic factors, the breast tumor microenvironment contributes to adaptive resistance [3]. Cancer-associated fibroblasts (CAFs) significantly impact tumor biology and are known to induce resistance to targeted therapies [4–9]. Despite this, CAFs are rarely incorporated into studies on breast cancer recurrence and resistance.
Patient-derived organoids (PDOs) are 3-dimensional multicellular clusters that are grown in basement membrane extract (BME). PDOs have self-renewal and self-organization capabilities and retain several features of the original tumor such as morphology and mutational landscape [10]. Compared to the traditional 2-dimensional cell cultures, PDOs capture intratumoral heterogeneity and are superior at predicting drug responses. To date, PDOs have been generated from several tumors, including breast cancer [11–13]. However, growing HR+ PDOs has been a challenge due to a low take rate (10%), overgrowth of normal breast epithelial cells, and the PDOs losing their HR expression and ceasing to grow after only a few passages [14,15]. In addition to PDOs, breast tumor organoids can be propagated from patient-derived xenografts (PDxO) [16,17]. Similar to the HR+ PDOs, HR+ PDxOs have a take rate of 9% for primary tumors and 16% for metastatic tumors. Currently, there are only a few models of metastatic HR+ breast cancer available [16,18]. Thus, there is an unmet need for novel models of HR+ primary and metastatic breast cancer that would advance our understanding of HR+ disease relapse and drug resistance.
We set out to develop patient-derived model systems for HR+ breast cancer that would also incorporate elements of the tumor microenvironment. Our optimized protocol allows for the efficient generation of HR+ PDOs from both primary and metastatic HR+ breast cancer with ~50% take rate. Importantly, we further optimized the media to support isolation and long-term passaging of HR+ bone metastasis that has been extremely difficult to grow. Our method also provides the advantage of reducing the cost and time by circumventing the necessity to passage through a mouse host. Furthermore, we isolate the matching CAFs that most protocols discard; thus, our optimized protocol offers the advantage of enabling the investigation of resistance mechanisms induced by the CAFs. Besides utilizing the patient-derived model for studying HR+ breast cancer and drug resistance [19–21], this model system can be applied to studying various aspects of tumor-stroma interactions.
Materials and reagents
Biological materials
1. Core needle biopsy, primary or metastatic breast cancer
2. Fine-needle aspiration (FNA), lymph node positive for tumor cells
3. Ascites, pleural, or peritoneal fluid
Reagents
1. Phosphate buffered saline (PBS) (Cytiva, catalog number: SH30028.FS)
2. Penicillin-streptomycin (Pen/Strep), 100× (Thermo Fisher, catalog number: 15140122)
3. Dispase-II solution (Sigma, catalog number: SCM133)
4. Collagenase type I (Sigma, catalog number: SCR103)
5. Human fibronectin solution (Sigma, catalog number: F0556)
6. Glucose solution, 200 g/L (Thermo Fisher, catalog number: A24940-01)
7. L-Glutamine solution, 200× (Thermo Fisher, catalog number: 25030-081)
8. HEPES solution, 1M (Sigma, catalog number: H0887-100ML)
9. Y-27632 2HCl, 10 mg (Selleckchem, catalog number: S1049)
10. Advanced DMEM/F12, 1× (Gibco, catalog number: 12634-010)
11. Primocin, 500 mg (InvivoGen, catalog number: ant-pm-05)
12. B27 supplement, 50× (Thermo Fisher, catalog number: 17504044)
13. Nicotinamide (Sigma, catalog number: N0636)
14. N-Acetyl-L-Cysteine (Sigma, catalog number: A9165)
15. A82-01 (Tocris, catalog number: 2939)
16. SB202190 (Selleckchem, catalog number: S1077)
17. 17β-Estradiol (Sigma, catalog number: 3301)
18. Hydrocortisone (Sigma, catalog number: H0888)
19. Human EGF (Thermo Fisher, catalog number: AF-100-15)
20. Human R-Spondin3 (Thermo Fisher, catalog number: 120-44)
21. Human noggin (Thermo Fisher, catalog number: 120-10C)
22. Human heregulin-β1 (Thermo Fisher, catalog number: 100-03)
23. Human FGF7 (Thermo Fisher, catalog number: 100-19)
24. Human FGF10 (Thermo Fisher, catalog number: 100-26)
25. Human CXCL12 (Thermo Fisher, catalog number: 300-28A)
26. Human IGF-I (Thermo Fisher, catalog number: 100-11)
27. Human osteopontin (Thermo Fisher, catalog number: 120-35)
28. Cultrex growth factor–reduced basement membrane extract type II (Fisher Scientific, catalog number: 353301002P)
29. Human fibroblast expansion basal medium, 1× (Thermo Fisher, catalog number: M106500)
30. Low serum growth supplement, 50× (Thermo Fisher, catalog number: S00310)
31. Fetal bovine serum (FBS) (R&D Systems, catalog number: S11550H)
32. Trypsin-EDTA, 0.25% (Thermo Fisher, catalog number: 25200056)
33. TrypLE Express, 1× (Thermo Fisher, catalog number: 12604013)
34. Dimethyl sulfoxide (DMSO) (Sigma, catalog number: D2650)
35. Corning cell recovery solution (Corning, catalog number: 354253)
36. CryoStor CS10 cryopreservation media (Sigma, catalog number: C2874)
37. Paraformaldehyde (PFA), 16%, EM grade (Electron Microscopy Sciences, catalog number: 15710-S)
38. HistoGelTM (Fisher Scientific, catalog number: 22-110-678)
39. Hematoxylin (Sigma, catalog number: MHS32-1L)
40. 10% formalin (Electron Microscopy Sciences, catalog number: 15740-04)
41. Bovine serum albumin (BSA) (Sigma, catalog number: A8806)
42. Goat serum (Thermo Fisher, catalog number: 16210064)
43. Triton X-100 (Sigma, catalog number: X100-500ML)
44. Cytoseal 60 (Fisher Scientific, catalog number: 23-244256)
45. Vectashield antifade mounting medium with DAPI (Fisher Scientific, catalog number: NC9524612)
46. Primary antibodies (Table 1)
Table 1. Primary antibodies for validation of ER status and characterization of CAFs
Antibody Host Company Catalog number
Estrogen receptor rabbit Abcam ab16666
Alexa 488 anti-rabbit secondary antibody goat Thermo Fisher A-11008
Fibronectin 1 (FN1) mouse Thermo Fisher MA5-1198
Platelet-derived growth factor receptor α (PDGFRα) rabbit Cell Signaling Technology 3174
Vimentin (VIM) rabbit Cell Signaling Technology 5741
α-Smooth muscle actin (αSMA) rabbit Abcam ab5694
Caveolin 1 (CAV1) rabbit Cell Signaling Technology 3267
Fibroblast activating protein (FAP) rabbit Cell Signaling Technology 66562
Podoplanin (PDPN) rabbit Cell Signaling Technology 9047
Thy1 cell surface antigen (THY1) rabbit Cell Signaling Technology 13801
Solutions
1. Biopsy collection solution (see Recipes)
2. Digestion solution (see Recipes)
3. PDO isolation wash solution (see Recipes)
4. Fibronectin coating solution (see Recipes)
5. Breast PDO media (see Recipes)
6. Bone metastasis PDO media (see Recipes)
7. Fibroblast media (see Recipes)
8. Freezing media (see Recipes)
9. IF blocking buffer (see Recipes)
10. IF wash buffer (see Recipes)
Recipes
1. Biopsy collection solution (500 mL)
Reagent Final concentration Quantity or Volume
PBS n/a 500 mL
Pen/Strep 100× 1× 5 mL
Glucose (200 g/L) 4.5 g/L 11.25 mL
2. Digestion solution (10 mL)
Make fresh for every digestion. Sterile filter before use.
Reagent Final concentration Quantity or Volume
Dispase-II solution 1× n/a 10 mL
Collagenase-I (200 mg/mL) 2 mg/mL 100 μL
Y-27632 2HCl (5 mM) 5 μM 10 μL
3. PDO wash solution (500 mL)
Reagent Final concentration Quantity or Volume
Advanced DMEM/F12 n/a 500 mL
Glutamine (200 mM) 2 mM 5 mL
HEPES (1 M) 10 mM 5 mL
Pen/Strep (100×) 1× 5 mL
Primocin (50 mg/mL) 50 μg/mL 500 μL
4. Fibronectin coating solution (5 mL)
Reagent Final concentration Quantity or Volume
PBS n/a 5 mL
Fibronectin 500 μg/mL 2 μg/mL 20 μL
5. Breast PDO media (50 mL)
Store at 4 °C for a maximum of two weeks. Add Y-27632 2HCl only for the first 4–6 days after isolation.
Reagent Final concentration Quantity or Volume
Advanced DMEM/F12 n/a 48 mL
Glutamine (200 mM) 2 mM 500 μL
HEPES (1 M) 10 mM 500 μL
Pen/Strep (100×) 1× 500 μL
Primocin (50 mg/mL) 50 μg/mL 50 μL
B27 supplement (50×) 1× 100 μL
Nicotinamide (1 M) 1 mM 50 μL
N-Acetyl-L-Cysteine (500 mM) 500 μM 50 μL
A82-01 (5 mM) 500 nM 5 μL
SB202190 (5 mM) 500 nM 5 μL
17β-Estradiol (5 μg/mL) 0.5 ng/mL 5 μL
Hydrocortisone (50 μg/mL) 50 ng/mL 50 μL
Human EGF (5 μg/mL) 5 ng/mL 50 μL
Human R-Spondin3 (100 μg/mL) 200 ng/mL 100 μL
Human noggin (40 μg/mL) 80 ng/mL 200 μL
Human heregulin-β1 (25 μg/mL) 25 ng/mL 50 μL
Human FGF7 (10 μg/mL) 5 ng/mL 25 μL
Human FGF10 (20 μg/mL) 20 ng/mL 50 μL
Y-27632 2HCl (5 mM) 5 μM 10 μL
6. Bone metastasis PDO media (50 mL)
Store at 4 °C for a maximum of two weeks. Add Y-27632 2HCl only for the first 4–6 days after isolation.
Reagent Final concentration Quantity or Volume
Advanced DMEM/F12 n/a 48 mL
Glutamine (200 mM) 2 mM 500 μL
HEPES (1 M) 10 mM 500 μL
Pen/Strep (100×) 1× 500 μL
Primocin (50 mg/mL) 50 μg/mL 50 μL
B27 supplement (50×) 1× 100 μL
Nicotinamide (1 M) 1 mM 50 μL
N-Acetyl-L-Cysteine (500 mM) 500 μM 50 μL
A82-01 (5 mM) 500 nM 5 μL
SB202190 (5 mM) 500 nM 5 μL
17β-Estradiol (5 μg/mL) 0.5 ng/mL 50 μL
Hydrocortisone (50 μg/mL) 50 ng/mL 50 μL
Human recombinant EGF (5 μg/mL) 5 ng/mL 50 μL
Human R-Spondin3 (100 μg/mL) 200 ng/mL 100 μL
Human noggin (40 μg/mL) 80 ng/mL 200 μL
Human heregulin-β1 (25 μg/mL) 25 ng/mL 50 μL
Human FGF7 (10 μg/mL) 5 ng/mL 25 μL
Human FGF10 (20 μg/mL) 20 ng/mL 50 μL
Human CXCL12 (10 μg/mL) 10 ng/mL 50 μL
Human IGF-I (20 μg/mL) 20 ng/mL 50 μL
Human osteopontin (10 μg/mL) 10 ng/mL 50 μL
Y-27632 2HCl (5 mM) 5 μM 10 μL
7. Fibroblast media (500 mL)
Reagent Final concentration Quantity or Volume
Human fibroblast expansion basal medium 1× n/a 500 mL
Low serum growth supplement 50× 1× 10 mL
FBS 4% 10 mL
Pen/Strep 100× 1× 5 mL
Primocin 50 mg/mL 50 μg/mL 500 μL
8. Freezing media (5 mL)
Reagent Final concentration Quantity or Volume
CryoStor CS10 n/a 5 mL
Y-27632 2HCl (5 mM) 5 μM 5 μL
9. IF blocking buffer (100 mL)
Reagent Final concentration Quantity or Volume
PBS n/a 100 mL
BSA 1% 1 g
Goat serum 5% 5 mL
Triton X-100 0.3% 300 μL
10. IF wash buffer (1,000 mL)
Reagent Final concentration Quantity or Volume
PBS n/a 1,000 mL
Triton X-100 0.3% 3 mL
Laboratory supplies
1. 50 mL centrifuge tube (Celltreat, catalog number: 229106)
2. 15 mL centrifuge tube (Celltreat, catalog number: 229411)
3. 25 mL serological pipettes, individually wrapped (Thermo Fisher, catalog number: 170357N)
4. 10 mL serological pipettes, individually wrapped (Thermo Fisher, catalog number:170356N)
5. 1,000 μL pipette tips (VWR, catalog number: 89082-350)
6. 200 μL pipette tips (VWR, catalog number: 89082-366)
7. 20 μL pipette tips (VWR, catalog number 89082-338)
8. 24-well plates (Thermo Fisher, catalog number: 142475)
9. 6-well plates (Thermo Fisher, catalog number: 140675)
10. 10 cm plates (Thermo Fisher, catalog number: 150350)
11. 1.5 mL microcentrifuge tubes (Eppendorf, catalog number: 0030120086)
12. 1.2 mL cryogenic vials (Corning, catalog number: 430487)
13. Mr. FrostyTM freezing container (Thermo Fisher, catalog number: 5100-0050)
14. TissueTek cryomold (VWR, catalog number: 25608-922)
15. #10 Bard-ParkerTM protected disposable scalpel (Fisher Scientific, catalog number: 02-688-78)
16. #11 Bard-ParkerTM protected disposable scalpel (Fisher Scientific, catalog number: 02-688-79)
17. Tissue cassette (VWR, catalog number: 18000-134)
18. BD disposable syringes (Fisher Scientific, catalog number: 14-823-435)
19. 25 G precision glide needles (Fisher Scientific, catalog number: 14-826-49)
20. FalconTM 4-well chamber slides (Fisher Scientific, catalog number: 08-774-209)
21. Coverglass 24 × 60 (Fisher Scientific, catalog number: NC1672857)
22. Forceps (Fisher Scientific, catalog number: 08-953G)
Equipment
1. Laminar flow hood (Baker, model: SterilGARD)
2. Incubator (Thermo, model: Forma Stericycle i160)
3. Centrifuge (Thermo, model: Sorvall legend XI)
4. Cell counter (Countess II FL)
5. Fisherbrand mini-tube rotator (Thermo Fisher, catalog number: 88-861-051)
6. Thermal mixer (Thermo Fisher, catalog number: 13687712)
7. Microscope (Olympus, model: CK2)
8. Microscope (Olympus, model: BX43)
9. Confocal microscope (Zeiss, model: LSM880)
Software and datasets
1. Zeiss (ZEN lite)
2. Fiji (version 2.14.0/1.54f)
Procedure
A. Isolation of patient-derived organoids from core-needle biopsies
1. Collect core-needle biopsies in a 50 mL tube with 10–20 mL of cold biopsy collection solution. Place tube on ice and transport to research laboratory within 30 min.
2. Use forceps to place one biopsy on a 6 cm dish with approximately 500 μL of digestion solution (Figure 1A).
Optional: Remaining biopsies can be placed in a cryotube with freezing media and frozen down and/or fixed using 4% PFA and used for histology. Frozen fragments can be stored at -196 °C for years.
Figure 1. Key steps during the isolation process. A) Place the core-needle biopsy into 500 μL of digestion solution. B) Slice the biopsy into 1–3 mm pieces. C) Move pieces into a 50 mL tube containing 10 mL of digestion solution. D) Stop digestion when pieces appear smaller and/or the digestion solution becomes “cloudy.” E) Spin down supernatant to pellet fragments. F) Pipette 50 μL of Cultrex and fragment mix into the middle of the well. G) Add PDO media by pipetting to the wall of the well.
3. Slice the biopsy into 1–3 mm pieces by using forceps and a 10-blade scalpel.
Note: Scissors can be used instead of a scalpel (Figure 1B).
4. Use forceps to move larger pieces to a 50 mL tube containing digestion solution. Pipette 1 mL of digestion solution to the 6 cm dish and move the solution containing smaller pieces to the 50 mL tube (Figure 1C).
5. Incubate biopsy pieces in an orbital shaker at 220 rpm for 45–60 min at 37 °C. Check on the progression of the digestion every 15 min. Stop the digestion when the pieces are visibly smaller or difficult to detect by the eye, and/or the digestion solution becomes “cloudy” (Figure 1D).
Critical: Do not exceed 60 min.
6. Shear the pieces by vigorously pipetting up and down using a 10 mL serological pipette. Let the fragments settle to the bottom of the tube and move the digestion solution to a new 50 mL tube. Add 10 mL of PDO wash solution supplemented with 5% FBS and pipette the pieces again. Move the wash solution containing tissue fragments to the same 50 mL tube.
7. Spin fragments down at 400× g for 5 min. Remove the supernatant and resuspend using 10 mL of fresh PDO isolation wash buffer (no FBS). Pipette 10 μL onto a glass slide and check for tissue fragments under a microscope. Spin down at 400× g for 5 min (Figure 1E).
Note: Pelleting the tissue fragments can be challenging if the biopsy has a high content of connective tissue, fat, and/or extracellular matrix. We transfer the supernatant into a separate 50 mL tube and store it until we have confirmed that we have obtained tissue fragments.
8. Remove supernatant and resuspend fragments in 1 mL of PDO isolation wash buffer. Count fragments using a hemocytometer and microscope (Olympus CK2). Set aside approximately 200 μL of the suspension for CAF isolation (section C). Pellet the fragments at 400× g for 5 min.
9. Embed fragments into Cultrex.
a. Thaw Cultrex on ice and keep it on ice for the whole duration.
b. Calculate the volume of Cultrex needed for the total number of fragments. Embed 800–1,000 fragments per 50 μL of Cultrex.
c. Carefully aspirate supernatant and resuspend pellet in Cultrex by slowly pipetting up and down using a 200 μL tip. This step should be done on ice.
d. Pipette 50 μL of the mix into the middle of the well on a 24-well plate or a 4-well chamber slide. The 50 μL drop should form a dome in the middle of the well (Figure 1F).
e. Place the 24-well plate into the incubator with standard conditions and incubate for 30 min at 37 °C.
10. Add 500 μL of breast PDO or bone metastasis PDO media supplemented with 5 μM Y-27632 2HCl to each well (Figure 1G). Change media every 4–5 days.
Note: Avoid adding media directly on top of the Cultrex dome.
11. Expand PDO cultures over the next 3–6 weeks. Image (Figure 2A) and freeze down PDOs during each passage.
a. Collect PDOs from 2–3 wells by scraping the bottom of the well with a 1,000 μL pipette tip and move the media and PDOs to a 15 mL tube. Spin down at 350× g for 5 min.
b. Resuspend the pellet with freezing media, move it to cryogenic tubes, and freeze using a Mr. FrostyTM freezing container.
Figure 2. Patient-derived organoids (PDO) morphology at different passages. A) Representative images of PDOs established from a core-needle biopsy. B) Representative images of PDOs established from ascites. Scale bar: 200 μm.
12. Validate ER expression after three passages by immunohistochemistry (see section E) or immunofluorescence staining (see section F).
B. Propagation of patient-derived organoids from fine-needle aspiration or ascites
1. Place the tube or bottle containing pleural/peritoneal fluid or FNA on ice and transport it to the research laboratory.
2. Spin down the fluid for 5 min at 400× g and aspirate supernatant. Repeat this step until all fluid has been pelleted.
3. Wash the pellet with PDO isolation wash solution and spin down for 5 min at 400× g. Repeat this step twice.
4. Resuspend the pellet in 1 mL of PDO isolation wash solution and calculate the cell number. Set aside approximately 200 μL of the suspension for CAF isolation (section C).
5. Embed cells into Cultrex.
a. Thaw Cultrex on ice and keep it on ice for the whole duration.
b. Calculate the volume of Cultrex needed for the total number of fragments. Embed 800–1,000 fragments per 50 μL of Cultrex.
c. Carefully aspirate supernatant and resuspend pellet in Cultrex by slowly pipetting up and down using a 200 μL tip. This step should be done on ice.
d. Pipette 50 μL of the mix into the middle of the well on a 24-well plate. The 50 μL drop should form a dome in the middle of the well (Figure 1F).
e. Place the 24-well plate into the incubator and incubate for 30 min.
6. Add 500 μL of breast PDO media Y-27632 2HCl to each well (Figure 1G). Change media every 4–5 days.
7. Expand PDO cultures over the next 3–6 weeks. Image (Figure 2B) and freeze down PDOs during each passage.
a. Collect PDOs from 2–3 wells by scraping the bottom of the well with a 1,000 μL pipette tip and move the media and PDOs to a 15 mL tube. Spin down for 5 min at 350× g.
b. Resuspend the pellet with freezing media, move it to cryogenic tubes, and freeze using a Mr. FrostyTM freezing container.
8. Validate ER expression after three passages by immunohistochemistry (see section E) or immunofluorescence staining (see section F).
C. Isolation and selection of cancer-associated fibroblasts
1. Add 1 mL of fibronectin coating solution per well to a 6-well plate and incubate the plate at 37 °C for 60 min. Wash wells with PBS before plating CAFs. We usually coat 2–3 wells.
2. Plate cell fragments and/or cells from step A8/B4 (Figure 1E) and add 2 mL of fibroblast media.
3. Enrich for CAFs by short trypsinization. CAFs proliferate faster than cancer cells and can be easily distinguished based on the elongated morphology. We usually observe cancer cells in 5%–10% of the initial CAF cultures (Figure 3). In case both CAFs and cancer cells are observed, perform a short trypsinization that will detach CAFs but leave cancer cells attached.
a. Aspirate media, wash cells with 4 mL of PBS, and add 1 mL of Trypsin-EDTA.
b. Incubate at 37 °C for a maximum of 5 min. Check on the cells every minute. Once cells start to detach, gently move the trypsin-containing CAFs into a 15 mL tube with 5 mL of fibroblast media.
Critical: Do not pipette or tap the plate to aid detachment.
c. Spin down at 350× g for 5 min and plate 1 × 106 cells/6 cm plate.
Figure 3. Representative images of a mixed population and a pure cancer-associated fibroblast (CAF) population. Scale bar: 200 μm.
4. Expand the CAFs over the next 3–6 weeks and freeze down early passages.
a. Wash plates with PBS, add 1.5 mL of trypsin-EDTA, and incubate for 5 min at 37 °C.
Note: Some CAF lines secret high amounts of extracellular matrix, which appears as a viscous membrane. We usually extend the incubation time with trypsin to allow complete detachment.
b. Add 6 mL of fibroblast media, move the suspension to a 15 mL tube, and pellet the CAFs at 350× g for 5 min.
c. Resuspend the pellet with freezing solution, move it to cryogenic tubes, and freeze using a Mr. FrostyTM freezing container. CAFs can be stored for years at -196 °C.
5. Characterize the CAFs by performing standard western blotting using the following fibroblast markers: FN1 (1:1,000), PDGFRα (1:1,000), αSMA (1:1,000), VIM1 (1:1,000), FAP, (1:1,000), THY1 (1:1,000), and PDPN (1:1,000).
D. Set up co-culture model
1. Collect and pellet PDOs from several wells by scraping the bottom of the well with a 1,000 μL pipette tip and move the media and PDOs to a 15 mL tube. Spin down for 5 min at 350× g.
2. Carefully aspirate the supernatant, add digestion solution, and incubate at 37 °C for 30–45 min. We add 500 μL of digestion solution per collected well.
Note: The pellet containing PDOs and Cultrex detaches easily from the bottom of the tube. We usually use a 1,000 μL pipette to remove the top layer of Cultrex.
3. Add 1 mL of PDO wash buffer, shear the PDOs by passing through a 25 G needle 10–20 times, and add 10 mL of PDO wash solution supplemented with 5% FBS. Some PDO lines are more difficult to break down into smaller fragments and require a 5 min TrypLE Express treatment. We advise to always passage new PDO lines with the digestion solution. Spin down for 5 min at 350× g, add 1 mL of PDO wash solution, and count the fragments.
4. Embed 200–600 fragments/well in the same way as in steps A9 and A10. Set up PDO cultures on a 24-well plate if the endpoint is a biochemical assay such as RNA isolation, or Falcon 4-well chamber slides for immunostainings (Figure 4).
Figure 4. Key steps for setting up a patient-derived organoid–cancer-associated fibroblast (PDO-CAF) co-culture
5. Set up co-culture 24 h later. Aspirate media, wash CAFs with PBS, and add 2 mL of trypsin to a 10 cm plate. Incubate for 5 min at 37 °C, add fibroblast media, and pellet cells in a 15 mL tube for 5 min at 350× g. Aspirate supernatant, resuspend pellet in 1 mL of fibroblast media, and count the cells.
6. Add 2,000–6,000 CAFs per well. The CAF number depends on how fast the CAF line proliferates.
a. Resuspend the correct amount of CAFs in breast PDO media or bone metastasis PDO media.
b. Pipette 500 μL of the suspension around the PDO dome.
c. Wait 24 h before starting any experiments with the co-cultures.
d. The length of the co-culture depends on the downstream application. We assess drug resistance for 4–5 days.
E. Embed PDOs in HistoGel
1. Collect PDOs from two wells by scraping the bottom of the well with a 1,000 μL pipette tip and move the media and PDOs to a 15 mL tube. Spin down for 5 min at 350× g.
2. Aspirate media, resuspend pellet in 1,000 μL of Corning cell recovery solution, and transfer the PDOs to a microcentrifuge tube.
3. Extract PDOs by incubating for 45–60 min at 4 °C on a rotor.
4. Set up inserts while PDOs are on the rotor.
a. Preheat HistoGel at 65 °C.
b. Cut the tip of a 200 μL tip and carefully add 200 μL of HistoGel to a cryomold. Avoid air bubbles.
c. Place the insert on ice.
5. Spin down PDOs for 5 min at 350× g, aspirate the supernatant, and fix PDOs in 500 μL of 4% PFA for 30 min at room temperature.
6. Add 1,000 μL of PBS, spin down for 5 min at 350× g, and aspirate the supernatant.
7. Stain PDOs using hematoxylin.
Note: This step is optional. We recommend staining the PDOs with hematoxylin as it makes the PDOs visible and easier to complete the remaining steps.
a. Gently resuspend pellet in 100 μL of hematoxylin and incubate for 5 min.
b. Add 1,000 μL of PBS, spin down for 5 min at 350× g, and carefully aspirate the supernatant.
8. Embed PDOs in HistoGel.
a. Cut the tip of a 200 μL tip and resuspend PDOs in 200 μL of preheated HistoGel.
b. Pipette the HistoGel and PDO mix on top of the cooled-down HistoGel layer from step F4.
c. Place cryomold on ice and incubate for at least 20 min.
d. Use an 11-blade scalpel to cut the HistoGel along the cryomold walls, invert the cryomold, and remove the HistoGel by gently pressing at the top of the cryomold.
e. Place HistoGel with PDOs into a tissue cassette.
9. Fix PDOs overnight at room temperature in 10% formalin and wash with PBS.
10. Embed in paraffin using standard protocol and stain for ER (1:100). Image using an Olympus BX43 microscope or similar.
F. Validation of ER expression by immunofluorescence staining
1. Aspirate media and fix PDOs in 500 μL of 4% PFA for 30 min at room temperature.
2. Gently remove PFA using a 1,000 μL pipette.
Note: The Cultrex dome becomes fragile and may spread toward the walls of the well during fixation. We usually aspirate the media from one corner of the well.
3. Wash with 500 μL of PBS. Repeat two times.
4. Permeabilize and block for 1 h at room temperature using 500 μL of blocking buffer.
5. Dilute ER antibody in blocking buffer at 1:100 ratio and incubate overnight at 4 °C.
6. Wash 10 times with IF wash buffer.
7. Dilute goat anti-rabbit secondary antibody 1:500 in blocking buffer and incubate for 2 h at RT.
Critical: Alexa 488 secondary antibody is light sensitive. Keep the culture slide covered from this step onward.
8. Wash 10 times with IF wash buffer.
9. Mount culture slide.
a. Wash once with H2O and carefully remove all H2O.
b. Remove the wells using the tool included with the culture slides.
c. Add Cytoseal 60 around the outer border of the culture slides.
d. Add a drop of Vectashield per well.
e. Gently press a coverslip on top of the culture slide. Avoid air bubbles.
f. Seal the staining by adding Cytoseal 60 around the outer border of the coverslip.
10. Image using a confocal microscope (Figure 5). We use a Zeiss LSM880 confocal microscope and Zeiss Zen Lite to convert acquired czi images into TIF format. Zeiss Zen Lite is the free version of Zen for basic image analysis.
Figure 5. Validation of estrogen receptor (ER) expression in patient-derived organoids (PDOs). Representative confocal image of ER- and DAPI-stained PDO. Scale bar: 40 μm.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Hogstrom et al. [20]. Simultaneous isolation of hormone receptor–positive breast cancer organoids and fibroblasts reveals stroma-mediated resistance mechanisms. Journal of Biological Chemistry [(Figure 1; Figure S1; Figure 2, panel A, B, D; Figure S2; Figure 3, panel B; Figure S3; Figure 5, panel B, C; Figure S5, panel A).
Hillis et al. [21]. Targeting Cholesterol Biosynthesis with Statins Synergizes with AKT Inhibitors in Triple-Negative Breast Cancer. Cancer Research. [(Figure 3, panel E–F; Supplementary figure S7)]
Choi et al. [19]. Lineage-specific silencing of PSAT1 induces serine auxotrophy and sensitivity to dietary serine starvation in luminal breast tumors. Cell Reports. [Figure 4, panel A, B, C, D).
General notes and troubleshooting
General notes
1. Our media recipe will support the growth of normal breast tissue. Similar to other published protocols, PDO cultures that are generated from primary tumors may be contaminated with normal breast organoids. It is important to validate ER expression and histology after three passages.
2. We recommend processing one biopsy and freezing down any additional biopsies as a backup.
3. Establishing and expanding PDOs and CAFs takes approximately 3–6 weeks. We set up co-cultures for downstream analysis once we have validated ER expression in PDOs and characterized CAFs with fibroblast markers. This usually takes place 4–8 weeks after establishing new PDO and CAF lines.
4. CAFs can be passaged 10–20 times before they undergo senescence. We recommend freezing as many vials as possible.
Troubleshooting
Problem 1: See General notes, point 1, regarding normal tissue contaminating the tumor PDOs.
Possible cause: The media supports the growth of normal breast tissue.
Solution: Validate the PDOs for tumor-specific markers, such as ER staining.
Problem 2: See General notes, point 3, regarding CAF undergoing senescence.
Possible cause: It is normal for CAFs to undergo senescence as they are not tumorigenic.
Solution: CAFs can be immortalized using hTERT to maintain replicative capacity beyond 20 passages. However, they will not be considered normal CAFs anymore at this point.
Acknowledgments
We thank all authors from our corresponding original research paper in the Journal of Biological Chemistry (2023) [20]. This research was supported by Susan G. Komen Foundation grant# CCR18547665, Harvard Stem Cell Institute grant # DP-0194-21-00, NIH/NCI grants #R00CA180221 and R21CA292302-01, and Ludwig Center at Harvard grant to T.M., J.M.H. was supported by Sigrid Juselius Foundation, Orion Research Foundation, Maud Kuistila Memorial Foundation and AACR-AstraZeneca Breast Cancer Research fellowship grant# 23-40-12-HOGS.
Figure 1 and Figure 4 were created in BioRender, License identifier: Muranen, T. (2024) BioRender.com/w03f535 and Muranen, T. (2024) BioRender.com/l38q446.
Competing interests
The authors declare that there are no conflicts of interest regarding the publication of this paper.
Ethical considerations
All experiments with human subjects were included in the IRB protocol that was reviewed by Dana-Farber/Harvard Cancer Center Scientific Review Committee and Institutional Review Board and approved in 2017 (#17-627). Informed consent was obtained from all participants as per Federal Regulations (45 CFR 46), BIDMC IRB Guidelines, and requirements of HIPAA, and the studies abide by the Declaration of Helsinki principles.
References
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Raheem, F., Karikalan, S. A., Batalini, F., El Masry, A. and Mina, L. (2023). Metastatic ER+ Breast Cancer: Mechanisms of Resistance and Future Therapeutic Approaches. Int J Mol Sci. 24(22): 16198.
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Article Information
Publication history
Received: Sep 3, 2024
Accepted: Nov 3, 2024
Available online: Dec 5, 2024
Published: Jan 20, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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5,161 | https://bio-protocol.org/en/bpdetail?id=5161&type=0 | # Bio-Protocol Content
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Peer-reviewed
Protocol to Retrieve Unknown Flanking DNA Using Fork PCR for Genome Walking
HW Hongjing Wu
HP Hao Pan
HL Haixing Li
Published: Vol 15, Iss 2, Jan 20, 2025
DOI: 10.21769/BioProtoc.5161 Views: 1082
Reviewed by: Alba BlesaFernando A Gonzales-Zubiate Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Frontiers in Microbiology Sep 2023
Abstract
PCR-based genome walking is one of the prevalent techniques implemented to acquire unknown flanking genomic DNAs. The worth of genome walking includes but is not limited to cloning full-length genes, mining new genes, and discovering regulatory regions of genes. Therefore, this technique has advanced molecular biology and related fields. However, the PCR amplification specificity of this technique needs to be further improved. Here, a practical protocol based on fork PCR is proposed for genome walking. This PCR uses a fork primer set of three arbitrary primers to execute walking amplification task, where the primary fork primer mediates walking by partially annealing to an unknown flank, and the fork-like structure formed between the three primers participates in inhibiting non-target amplification. In primary fork PCR, the low-annealing temperature (25 °C) cycle allows the primary fork primer to anneal to many sites of the genome, synthesizing a cluster of single-stranded DNAs; the subsequent 65 °C cycle processes the target single-strand into double-strand via the site-specific primer; then, the remaining 65 °C cycles selectively enrich this target DNA. However, any non-target single-stranded DNA formed in the 25 °C cycle cannot be further processed in the following 65 °C cycles because it lacks an exact binding site for any primer. Secondary, or even tertiary nested fork PCR further selectively enriches the target DNA. The practicability of fork PCR was validated by walking three genes in Levilactobacillus brevis CD0817 and one gene in Oryza sativa. The results indicated that the proposed protocol can serve as a supplement to the existing genome walking protocols.
Key features
• This protocol builds upon the method developed by Pan et al. [1], which is applicable to genome-walking for any species.
• The developed protocol is a random priming PCR-based genome-walking scheme.
• Two rounds of nested fork PCR amplifications suffice to release a positive walking result.
Keywords: Genome walking Site-specific primer Fork primer Overlap of fork primers Partial annealing Intra-strand annealing Electrophoresis DNA sequencing
Graphical overview
Background
Genome walking refers to a strategy used to mine unknown genomic regions flanking known DNAs. Genome walking has promoted the development of molecular biology–related fields by being widely applied, such as in cloning full-length genes, uncovering transgenic sites, and acquiring regulatory regions of genes. To date, many PCR-based genome walking techniques have been developed [2–6]. These techniques fall into two clusters according to the involved methodological principles: cluster I, genome pre-processing-dependent PCR [7–10], and cluster II, random priming PCR [11–13].
A random priming PCR method omits the genome pre-processing prior to PCR, having thus attracted increasing interest from researchers [14–16]. Over the past decades, about a dozen random PCR methods have been constructed, such as wristwatch PCR [4,17], racket PCR [3,5], and thermal asymmetric interlaced PCR [18,19]. In such a PCR, an arbitrary primer mediates genome walking by partially hybridizing to an unknown flank genomic region in the low-temperature cycle, while a nested sequence-specific primer(s) (NSP) ensures PCR amplification specificity. In general, this method requires at least two rounds of nested amplifications to obtain a target amplicon(s) but still suffers from a non-target background arising from the arbitrary primer [4,14]. Therefore, developing a random PCR genome walking method with satisfactory amplification specificity remains of interest.
Recently, we have developed a new genome-walking technique called fork PCR. The fork PCR depends on the partial overlaps between the three arbitrary primers [primary fork primer (PFP), secondary fork primer (SFP), and branch primer (BP)] in a fork primer set. PFP and SFP share a 3' part (stem), while their 5' parts (branches) are heterologous to each other. BP corresponds to the 5' branch of SFP. Therefore, PFP, SFP, and BP form a fork-like structure (Figure 1). The fork PCR has shown satisfactory amplification specificity mainly due to the following two facts: First, all thermal cycles in secondary/tertiary fork PCR are stringent (annealing temperature 65 °C). Second, the non-target product defined by PFP is hard to amplify in secondary/tertiary PCR as it tends to form a hairpin via intra-strand annealing mediated by the inverted PFP terminal repeats, rather than being annealed by SFP or BP. The feasibility of fork PCR has been verified by walking several selected genetic sites [1].
Materials and reagents
Biological materials
1. Genomic DNA of Levilactobacillus brevis CD0817 [20–23], prepared by our lab at Nanchang University (Nanchang, China)
2. Genomic DNA of Oryza sativa, obtained from the Lab of Dr. Xiaojue Peng at Nanchang University (Nanchang, China)
Reagents
1. LA Taq polymerase (hot-start version) (Takara, catalog number: RR042A)
2. dNTP mixture (Takara, catalog number: RR042A)
3. 10× LA PCR buffer (Mg2+ plus) (Takara, catalog number: RR042A)
4. 6× Loading buffer (Takara, catalog number: 9156)
5. DL 5,000 DNA marker (Takara, catalog number: 3428Q)
6. 1× TE buffer (Sangon, catalog number: B548106)
7. Agarose (Sangon, catalog number: A620014)
8. 1 M NaOH (Yuanye, catalog number: B28412)
9. Green fluorescent nucleic acid dye (10,000×) (Solarbio, catalog number: G8140)
10. 0.5 M EDTA (Solarbio, catalog number: B540625)
11. Boric acid (Solarbio, catalog number: B8110)
12. Tris (Solarbio, catalog number: T8060)
13. DiaSpin DNA Gel Extraction kit (Sangon, catalog number: B110092)
14. Primers (Sangon)
PFP1: 5'-ACGCGTAATAGCTCGGGATGATGCTGCTCGTGGATGACTCT-3'
SFP1: 5'-CCTGACCGCCTTCTACACCTATGCTGCTCGTGGATGACTCT-3'
PFP2: 5'-ATCCGCCCATAGCCTTCAGTGACTACGCTGCCTTGCTACTT-3'
SFP2: 5'-CCTGACCGCCTTCTACACCTGACTACGCTGCCTTGCTACTT-3'
BP: 5'-CCTGACCGCCTTCTACACCT-3'
oNSP-gadA: 5'-GTTTCTGGTCACAAGTACGGCATGG-3'
mNSP-gadA: 5'-TGCTGATACGCTGCCAGAAGAAATG-3'
iNSP-gadA: 5'-ACGGTTGACTCCATTGCCATTAACT-3'
oNSP-gadR: 5'-TCCTTCGTTCTTGATTCCATACCCT-3'
mNSP-gadR: 5'-CCATTTCCATAGGTTGCTCCAAGG-3'
iNSP-gadR: 5'-GGATACTGGCTAAAATGAATTAACTCGGATAA-3'
oNSP-pct: 5'-TCTTGTTCTTCAACAGTGGTGGGTA-3'
mNSP-pct: 5'-TCGTCTTTCGTGTAAGTGTTGGTGT-3'
iNSP-pct: 5'-AGGAAATATGCACTCTTGGGAAGCG-3'
oNSP-hyg: 5'-ACGGCAATTTCGATGATGCAGCTTG-3'
mNSP-hyg: 5'-GGGACTGTCGGGCGTACACAA-3'
iNSP-hyg: 5'-CTGGACCGATGGCTGTGTAGAAG-3'
Solutions
1. 2.5× TBE buffer (see Recipes)
2. 0.5× TBE buffer (see Recipes)
3. 100 μM primer (see Recipes)
4. 10 μM primer (see Recipes)
5. 1% agarose gel (see Recipes)
Recipes
1. 2.5× TBE buffer
Reagent Final concentration Amount
0.5 M EDTA solution 5 mM 10 mL
Tris 225 mM 27 g
Boric acid 225 mM 13.75 g
ddH2O n/a 950 mL
Total n/a 1,000 mL
Adjust pH to 8.3 with 1 M NaOH and then top the solution to 1,000 mL with ddH2O.
2. 0.5× TBE buffer
Reagent Final concentration Amount
2.5× TBE buffer 0.5× 200 mL
ddH2O n/a 800 mL
Total n/a 1,000 mL
3. 100 μM primer
Reagent Final concentration Quantity or Volume
Powdery primer 100 μM n/a
1× TE buffer 1× Volume specified in the sheet of primer synthesis
Total n/a Volume specified in the sheet of primer synthesis
Note: Dilute a portion of the 100 μM primer to prepare 10 μM primer and store the remaining portion at -80 °C.
4. 10 μM primer
Reagent Final concentration Quantity or Volume
100μM primer 10 μM 1 μL
1× TE buffer 1× 9 μL
Total n/a 10 μL
Note: Prepare extra volume of a 10 μM primer and pipette it to multiple 1.5 mL microcentrifuge tubes. Then, store the tubes at -80 °C. Take one tube at a time and store it at -20 °C after use.
5. 1% agarose gel
Reagent Final concentration Quantity or Volume
Agarose 1% 1 g
0.5× TBE buffer 0.5× 100 mL
Green fluorescent nucleic acid dye (10,000×) 1× 10 μL
Total n/a 100 mL
Laboratory supplies
1. 0.2 mL thin-wall PCR tubes (Kirgen, catalog number: KG2311)
2. 10 μL pipette tips (Sangon, catalog number: F600215)
3. 200 μL pipette tips (Sangon, catalog number: F600227)
4. 1,000 μL pipette tips (Sangon, catalog number: F630101)
5. 1.5 mL microcentrifuge tubes (Labselect, catalog number: MCT-001-150)
Equipment
1. PCR apparatus (Analtytikjena, model: Biometra TAdvanced)
2. Microcentrifuge (Tiangen, model: TGear)
3. Electrophoresis apparatus (Beijing Liuyi, model: DYY-6C)
4. Gel imaging system (Bio-Rad, model: ChemiDoc XRS+)
Software and datasets
1. Oligo 7 software (Molecular Biology Insights, Inc., USA)
2. DNASTAR Lasergene software (DNASTAR, Inc.)
Procedure
A. Design of primers
1. Select three NSPs—outmost NSP (oNSP), middle NSP (mNSP), and innermost NSP (iNSP)—from a known DNA.
Critical: The Tm values of NSPs are from 60 to 65 °C. An NSP itself should avoid forming a hairpin structure with a Tm value exceeding 40 °C.
2. Design two sets of fork primers. The three primers (PFP, SFP, and BP) in each set form a fork-like structure (Figure 1).
Figure 1. Fork-like structures of the two walking primer sets. PFP or SFP consists of 5' branch and 3' stem. The stems of the two primers are homologous to each other, while the branches are heterologous to each other. BP corresponds to the branch of SFP. The melting temperatures of PFP1, SFP1, PFP2, SFP2, and BP are 73.6, 74.2, 74.6, 74.5, and 60.7 °C, respectively. BP is universal to the two fork primer sets. PFP: primary fork primer, SFP: secondary fork primer, and BP: branch primer.
Critical: The sequence of a fork primer (PFP, SFP, or BP) is completely arbitrary, with the four bases adenine (A), thymine (T), cytosine (C), and guanine (G) being evenly distributed. Meanwhile, a fork primer itself should avoid forming a severe hairpin or dimer structure and meet the criteria shown in Table 1. Simultaneously designing more than one fork primer set is suggested to execute parallel fork PCRs in a walking cycle.
Table 1. Key criteria for designing fork primer
Primer Length (nt) G+C content (%) Melting temperature (°C)
PFP 41 40–60 ~75
SFP 41 40–60 ~75
BP 20 40–60 60–65
PFP: primary fork primer; SFP: secondary fork primer; BP: branch primer; G: guanine; C: cytosine.
Note: Use the Oligo 7 software to devise and assess primers.
B. Fork PCR amplifications
A fork PCR set comprises three rounds of nested amplifications. Figure 2 describes the process of fork PCR.
Figure 2. Schematic diagram of fork PCR. oNSP: outmost nested site-specific primer; mNSP: middle nested site-specific primer; iNSP: innermost nested site-specific primer; PFP: primary fork primer; SFP: secondary fork primer; BP: branch primer; HSC: high-stringency cycle; LSC: low-stringency cycle. Thin solid line: known DNA; dotted line: unknown DNA; arrows: primers; thick solid lines: primer complements.
Critical: The working concentration of SFP is 10% of that of BP or mNSP.
Note: In secondary fork PCR, types I (defined by oNSP) and II (defined by oNSP and PFP) non-target products are readily removed because they lack an exact binding site for mNSP. Type III non-target product (defined by PFP) tends to form hairpin via the PFP termini rather than binding with SFP, because PFP has a higher Tm value than the overlap between PFP and SFP. Clearly, the amplification of type III non-target product is also inhibited. However, the target product can be exponentially enriched once SFP integrates into the PFP site. Tertiary fork PCR further selectively enriches the target product.
1. Primary fork PCR
a. Pipette primary PCR components (Table 2) into a 0.2 mL PCR tube.
Table 2. Primary fork PCR mix
Reagent Final concentration Amount (μL)
Genomic DNA Microbe, 0.2–2 ng/μL; plant or animal, 2–20 ng/μL 1
LA Taq polymerase (5 U/μL) 0.05 U/μL 0.5
PFP (10 μM) 0.2 μM 1
oNSP (10 μM) 0.2 μM 1
10× LA PCR buffer II (Mg2+ plus) 1× 5
dNTP mixture (2.5 mM each) 0.4 mM each 8
ddH2O n/a 33.5
Total n/a 50
b. Completely mix the components with a pipette.
c. Centrifuge for 10–20 s with a microcentrifuge to gather the mixture.
d. Run the amplification in the PCR apparatus (Table 3).
Table 3. Primary fork PCR cycling conditions
Step Temperature (°C) Duration Cycle
Initial denaturation 95 2 min 1
Denaturation 95 10 s 1
Annealing 25 30 s
Extension 72 2 min
Denaturation 95 10 s 30
Annealing 65 30 s
Extension 72 2 min
Final extension 72 5 min 1
Hold 4 forever
e. Put the PCR product onto ice.
f. Take 1 μL of the product as the template of secondary fork PCR.
g. Store the remaining product at -20 °C for future assays.
2. Secondary fork PCR
a. Pipette secondary PCR components (Table 4) into a 0.2 mL PCR tube.
Table 4. Secondary fork PCR mix
Reagent Final concentration Amount (μL)
Primary PCR product n/a 1
LA Taq polymerase (5 U/μL) 0.05 U/μL 0.5
BP (10 μM) 0.2 μM 1
SFP (1 μM) 0.02 μM 1
mNSP (10 μM) 0.2 μM 1
10× LA PCR buffer II (Mg2+ plus) 1× 5
dNTP mixture (2.5 mM each) 0.4 mM each 8
ddH2O n/a 32.5
Total n/a 50
Note: The working concentration of SFP is 10% of that of BP or mNSP.
Critical: Dilute primary PCR product 10–1,000 fold if necessary.
b. Completely mix the components with a pipette.
c. Centrifuge for 10–20 s with a microcentrifuge to gather the mixture.
d. Run the amplification in the PCR apparatus (Table 5).
Table 5. Secondary fork PCR cycling conditions
Step Temperature (°C) Duration Cycle
Initial denaturation 95 2 min 1
Denaturation 95 10 s 30
Annealing 65 30 s
Extension 72 2 min
Final extension 72 5 min 1
Hold 4 forever
e. Put the PCR product onto ice.
f. Take 1 μL of the product as the template of tertiary fork PCR.
g. Store the remaining product at -20 °C for future assays.
3. Tertiary fork PCR
a. Pipette tertiary amplification components (Table 6) into a 0.2 mL PCR tube.
Table 6. Tertiary fork PCR mix
Reagent Final concentration Amount (μL)
Secondary PCR product n/a 1
LA Taq polymerase (5 U/μL) 0.05 U/μL 0.5
BP (10 μM) 0.2 μM 1
iNSP (10 μM) 0.2 μM 1
10× LA PCR buffer II (Mg2+ plus) 1× 5
dNTP mixture (2.5 mM each) 0.4 mM each 8
ddH2O n/a 33.5
Total n/a 50
Critical: Dilute secondary PCR product 10–1,000 fold if necessary.
b. Completely mix the components with a pipette.
c. Centrifuge for 10–20 s with a microcentrifuge to gather the mixture.
d. Run the amplification in the PCR apparatus (Table 7).
Table 7. Tertiary fork PCR cycling conditions
Step Temperature (°C) Duration Cycle
Initial denaturation 95 2 min 1
Denaturation 95 10 s 30
Annealing 65 30 s
Extension 72 2 min
Final extension 72 5 min 1
Hold 4 forever
e. Store the PCR product at -20 °C for future assays.
C. Gel electrophoresis
1. Completely mix 5 μL of PCR product and 1 μL of 6× loading buffer.
2. Load the mixture into 1% agarose gel supplemented with 1× green fluorescent nucleic acid dye.
3. Set the electrophoresis apparatus to a voltage of 150 V (the distance between the two electrodes is 30 cm).
4. Check the PCR product using the ChemiDoc XRS+ imaging system after approximately 25 min of electrophoresis (Figure 3).
Figure 3. Fork PCR to genes gadA and hyg. Forks 1 and 2 denote the two parallel fork PCR sets in the walking experiment. The fragments marked with white arrows indicate the target products. Lanes P, S, and T show primary, secondary, and tertiary PCR, respectively. Lane M: TaKaRa DL5000 Marker.
D. Recovery of PCR product
1. Completely mix 40 μL of secondary/tertiary fork PCR product and 8 μL of 6× loading buffer.
2. Load the mixture into 1% agarose gel supplemented with 1× green fluorescent nucleic acid dye.
3. Set the electrophoresis apparatus to a voltage of 150 V (the distance between the two electrodes is 30 cm).
4. Visualize the PCR product using the ChemiDoc XRS+ imaging system. Subsequently, cut out clear DNA band(s) using a knife.
5. Extract DNA from the cut gel using the DiaSpin DNA Gel Extraction kit.
6. Confirm the extracted DNA with 1% agarose gel electrophoresis.
E. DNA sequencing
Directly sequence the extracted DNA at Sangon Biotech Co., Ltd.
Data analysis
The correctness of obtained DNA is tested by sequence alignment performed by the “By Clustal W Method” function in MegAlign software. The walking is regarded as correct if one end of the obtained DNA sequence overlaps with one end of the known DNA [24,25].
1. Open the MegAlign software, click File, and then click Enter Sequences (Figure 4).
Figure 4. Screenshot of the MegAlign software showing the location of the Enter Sequences under the File tab
2. Input a DNA sequence walked by fork PCR and the corresponding known DNA segment between iNSP and unknown flank.
3. Click Align, and then click the By Clustal W Method (Figure 5) to output the alignment result (Figure 6).
Figure 5. Screenshot of the MegAlign software displaying the input DNA sequences
Figure 6. Screenshot of the MegAlign software displaying the alignment result
Note: The walking is considered successful if the iNSP-sided part of the PCR product overlaps the known DNA.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article:
Pan et al. [1] Fork PCR: a universal and efficient genome-walking tool. Frontiers in Microbiology (Figure 3).
General notes and troubleshooting
General notes
1. Three primers, SFP, BP, and mNSP, are used in secondary fork PCR. However, the working concentration of SFP is only 10% of that of BP or mNSP. The major role of SFP is to integrate the BP sequence into the PFP site, and the real amplifiers are BP and mNSP.
2. Primary fork PCR product can be exempted from testing.
3. After secondary fork PCR, the PCR product is checked using agarose gel electrophoresis to determine whether a clear DNA band(s) appears. If a clear DNA band(s) appears, it is recovered and sequenced. There is no need to perform tertiary PCR if this DNA band(s) is correct.
4. Like the other PCR-based walking protocols, the current protocol also suffers from the multiple-band phenomenon. In general, only the largest DNA band needs to be analyzed if multiple DNA bands appear.
5. Simultaneously performing parallel fork PCRs in a walking cycle will improve walking success and efficiency.
6. The two fork primer sets provided here are universal to any species. A researcher needs only to design an NSP set in a walking experiment. An NSP itself should avoid forming a severe hairpin or dimer; in addition, it should avoid forming a severe dimer with the paired fork primer.
7. Researchers can also design more fork primer sets as they wish, provided that they meet the criteria mentioned above. N fork primer sets require n PFPs, n SFP, and one BP. The BP is universal to the N fork primer sets.
Troubleshooting
Problem 1: No clear DNA band(s) appears in secondary or even tertiary fork PCR.
Possible cause: In primary fork PCR, non-target amplification is very strong, while target amplification is very weak.
Solution: Dilute primary PCR product 10–1,000 times; then, use 1 μL dilutions as templates for secondary fork PCRs. Afterward, perform the corresponding tertiary PCRs. If still no clear DNA(s) appears in any secondary/tertiary PCR, redesign an NSP set with a different position from the previous one.
Problem 2: The multi-band phenomenon is serious, which affects the cutting and recovery of target PCR products from agarose gel.
Possible cause: In primary fork PCR, PFP randomly anneals to many sites on unknown flanking DNA.
Solution: Dilute the primary PCR product 10–1,000 times and use 1 μL dilutions as templates for secondary fork PCRs. Then, select a satisfactory secondary PCR. In fact, the multi-band phenomenon is common in random priming PCR-based genome-walking methods but, generally, only the largest DNA band needs to be considered.
Problem 3: Direct sequencing of fork PCR product is not smooth.
Possible cause: There is interference from the non-target background.
Solution: Clone the product and then sequence it.
Problem 4: Clear DNA band(s) is the non-target product.
Possible cause: Genomic DNA template may be contaminated.
Solution: Use an uncontaminated genomic DNA template, use dedicated sterile consumables, and perform PCR in a separate and clean region.
Acknowledgments
This study was supported by the Jiangxi Provincial Department of Science and Technology (Grant no. 20225BCJ22023), China and the National Natural Science Foundation of China (Grant no. 32160014), China. This fork PCR-based genome-walking protocol has been originally described and validated in Frontiers in Microbiology [1].
Competing interests
The authors declare no competing interests.
References
Pan, H., Guo, X., Pan, Z., Wang, R., Tian, B. and Li, H. (2023). Fork PCR: a universal and efficient genome-walking tool. Front Microbiol. 14: e1265580.
Kotik, M. (2009). Novel genes retrieved from environmental DNA by polymerase chain reaction: Current genome-walking techniques for future metagenome applications. J Biotechnol. 144(2): 75–82.
Sun, T., Jia, M., Wang, L., Li, Z., Lin, Z., Wei, C., Pei, J. and Li, H. (2022). DAR-PCR: a new tool for efficient retrieval of unknown flanking genomic DNA. AMB Express. 12(1): 131.
Wang, L., Jia, M., Li, Z., Liu, X., Sun, T., Pei, J., Wei, C., Lin, Z. and Li, H. (2022). Wristwatch PCR: A Versatile and Efficient Genome Walking Strategy. Front Bioeng Biotechnol. 10: e792848.
Pei, J., Sun, T., Wang, L., Pan, Z., Guo, X. and Li, H. (2022). Fusion primer driven racket PCR: A novel tool for genome walking. Front Genet. 13: e969840.
Wang, R., Gu, Y., Chen, H., Tian, B. and Li, H. (2025). Uracil base PCR implemented for reliable DNA walking. Anal Biochem. 696: 115697.
Yik, M. Y., Lo, Y. T., Lin, X., Sun, W., Chan, T. F. and Shaw, P. C. (2021). Authentication of Hedyotis products by adaptor ligation-mediated PCR and metabarcoding. J Pharm Biomed Anal. 196: 113920.
Uchiyama, T. and Watanabe, K. (2006). Improved inverse PCR scheme for metagenome walking. Biotechniques. 41(2): 183–188.
Alquezar‐Planas, D. E., Löber, U., Cui, P., Quedenau, C., Chen, W. and Greenwood, A. D. (2020). DNA sonication inverse PCR for genome scale analysis of uncharacterized flanking sequences. Methods Ecol Evol. 12(1): 182–195.
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Lin, Z., Wei, C., Pei, J. and Li, H. (2023). Bridging PCR: An efficient and reliable scheme implemented for genome-walking. Curr Issues Mol Biol. 45(1): 501–511.
Wei, C., Lin, Z., Pei, J., Pan, H. and Li, H. (2023). Semi-site-specific primer PCR: A simple but reliable genome-walking tool. Curr Issues Mol Biol. 45(1): 512–523.
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Wang, L., Jia, M., Li, Z., Liu, X., Sun, T., Pei, J., Wei, C., Lin, Z. and Li, H. (2023). Protocol to access unknown flanking DNA sequences using Wristwatch-PCR for genome-walking. STAR Protoc. 4(1): 102037.
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Article Information
Publication history
Received: Sep 14, 2024
Accepted: Nov 17, 2024
Available online: Nov 27, 2024
Published: Jan 20, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Cloning a Chloroplast Genome in Saccharomyces cerevisiae and Escherichia coli
EW Emma Jane Lougheed Walker §
BK Bogumil Jacek Karas §
(§ Technical contact)
Published: Vol 15, Iss 2, Jan 20, 2025
DOI: 10.21769/BioProtoc.5162 Views: 2047
Reviewed by: Lucy XieSean L. BeckwithEmilia Krypotou
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Original Research Article:
The authors used this protocol in Plant Physiology Apr 2024
Abstract
Chloroplast genomes present an alternative strategy for large-scale engineering of photosynthetic eukaryotes. Prior to our work, the chloroplast genomes of Chlamydomonas reinhardtii (204 kb) and Zea mays (140 kb) had been cloned using bacterial and yeast artificial chromosome (BAC/YAC) libraries, respectively. These methods lack design flexibility as they are reliant upon the random capture of genomic fragments during BAC/YAC library creation; additionally, both demonstrated a low efficiency (≤ 10%) for correct assembly of the genome in yeast. With this in mind, we sought to create a highly flexible and efficient approach for assembling the 117 kb chloroplast genome of Phaeodactylum tricornutum, a photosynthetic marine diatom. Our original article demonstrated a PCR-based approach for cloning the P. tricornutum chloroplast genome that had 90%–100% efficiency when screening as few as 10 yeast colonies following assembly. In this article, we will discuss this approach in greater depth as we believe this technique could be extrapolated to other species, particularly those with a similar chloroplast genome size and architecture.
Key features
• Large fragments of the chloroplast genome can be readily amplified through PCR from total algal DNA isolate.
• Assembly protocol can be completed within a day, and yeast colonies harboring chloroplast genomes can be obtained in as few as 4–5 days.
• Cloned genomes isolated from yeast transformants can be moved to Escherichia coli through electroporation.
Keywords: Synthetic biology Chloroplast Genome engineering Genome replacement Yeast assembly Cloning Transformation Phaeodactylum tricornutum Escherichia coli PCR
Graphical overview
The chloroplast genome is split into overlapping fragments (green), which are then PCR-amplified along with a suitable cloning vector (blue). The fragments are transformed into Saccharomyces cerevisiae for assembly, where they will recombine to form the cloned chloroplast genome. DNA can be isolated from yeast transformants and electroporated into Escherichia coli for further analysis. Ultimately, to confirm the capture of the cloned genome, DNA from representative yeast and/or E. coli transformants is sequenced.
Background
The ability to assemble, deliver, and install whole synthetic genomes provides the utmost control when engineering an organism. However, synthesizing large fragments of DNA (i.e., > 10 kb) is still prohibitively expensive for most academic labs, whilst cloning and transforming partial or whole chromosomes remains technically challenging, if not unexplored, in most species. The model systems for whole-genome replacement have been limited to comparatively simple organisms with well-established DNA manipulation techniques (e.g., viruses [1,2], bacteria [3,4], and Saccharomyces cerevisiae [5]). Given the exciting possibilities that whole-genome replacement offers, there is a growing demand to establish other biological chassis capable of large-scale engineering feats [6]. Establishing a photosynthetic chassis is of particular interest to the biotechnology and bioeconomy sectors, as photosynthetic organisms can capture and use atmospheric CO2 as a carbon source, forgoing the need for energetically costly carbon inputs.
Photosynthetic eukaryotes possess distinct nuclear, chloroplast, and mitochondrial genomes. The aforementioned organelles typically contain multiple copies of a single, highly reduced chromosome, making these genomes more feasible to synthesize and “replace” compared to nuclear chromosomes, which are larger and more complex. The chloroplast genome is a particularly interesting target for genome replacement as the organelle serves as a hub for several cellular biosynthesis pathways of industrial relevance (e.g., starch, amino acid, and fatty acid synthesis). Despite this, there are only two published papers demonstrating the potential for cloning an entire chloroplast genome outside of the organelle. The first paper describes the capture of the Zea mays chloroplast genome (140 kb) in Saccharomyces cerevisiae [7]. Here, high molecular weight DNA was isolated from Z. mays, then shorn and ligated into a yeast artificial chromosome (YAC) backbone. More than 10,000 yeast clones were screened, with only one clone demonstrating what appeared to be the complete Z. mays chloroplast genome. The second paper describes the assembly of the Chlamydomonas reinhardtii chloroplast genome (204 kb) from six overlapping fragments, of which four had been previously captured in a bacterial artificial chromosome (BAC) library [8]. Here, 3 out of 30 yeast transformants contained the correctly assembled chloroplast genome following yeast assembly. Creating BAC/YAC libraries is technically demanding, time consuming, and does not permit the utmost design flexibility for assembling large constructs as it is dependent upon whichever fragments were generated during the library preparation stage. An efficient and tractable strategy for assembling whole chloroplast genomes is a necessary first step to achieve the full potential of this unique chassis.
We sought to design and test alternative assembly strategies for cloning whole chloroplast genomes. Phaeodactylum tricornutum was selected as our model organism due to its ease of propagation and rapidly growing toolbox for genetic engineering. Here, we present a strategy for cloning the P. tricornutum chloroplast genome that demonstrated 90%–100% efficiency when screening as few as 10 yeast colonies and 3 Escherichia coli colonies following whole-genome assembly and transformation, respectively. We believe this efficient and tractable method for cloning the P. tricornutum chloroplast genome could be extrapolated to other photosynthetic eukaryotes, particularly those with similar genome sizes and architectures (e.g., Thalassiosira pseudonana).
Materials and reagents
Biological materials
1. P. tricornutum liquid culture [Culture Collection of Algae, the University of Texas at Austin (UTEX), catalog number: 646]
2. E. coli TransforMax EPI300 cells (LGC Biosearch Technologies, Lucigen, catalog number: EC300110)
3. S. cerevisiae VL6-48 culture [American Type Culture Collection (ATCC), catalog number: MYA-3666]
Reagents
1. 1 kb DNA ladder (New England Biolabs, catalog number: N3232L)
2. 2-Mercaptoethanol (Sigma-Aldrich, catalog number: M3148)
3. Adenine hemisulfate salt (Sigma-Aldrich, catalog number: A2545)
4. Agar A (Bio Basic, catalog number: FB0010)
5. Agarose (FroggaBio, catalog number: A87-500G)
6. Bacteriological peptone (BioShop, catalog number: PEP403)
7. Bacto agar (Becton Dickinson, catalog number: 214030)
8. Bio-tryptone (BioShop, catalog number: TRP402)
9. Boric acid (Bio Basic, catalog number: BB0044)
10. Calcium chloride, dihydrate (BioShop, catalog number: CCL302)
11. Cetyltrimethylammonium bromide (CTAB) (Sigma-Aldrich, catalog number: H6269)
12. Chloramphenicol (Bio Basic, catalog number: CB0118)
13. Chloroform:isoamyl alcohol, 24:1 (Bio Basic, catalog number: CB0351)
14. Cobalt (II) sulfate, heptahydrate (Sigma-Aldrich, catalog number: CDS004010)
15. Complete media glucose broth minus tryptophan (Teknova, catalog number: C7131)
Note: This product is no longer available; as an alternative, we suggest minimal SD base (Takara, catalog number: 630411) and -Trp DO supplement (Takara, catalog number: 630413).
16. Complete media glucose broth minus histidine and uracil (Teknova, catalog number: C7221)
Note: This product is no longer available; as an alternative, we suggest minimal SD base (Takara, catalog number: 630411) in addition to the minus histidine and uracil (-His/-Ura) DO supplement (Takara, catalog number: 630422).
17. Copper (II) sulfate, pentahydrate (Bio Basic, catalog number: CDB0063)
18. Cyanocobalamin, i.e., Vitamin B12 (BioShop, catalog number: VIT271)
19. D-biotin (Bio Basic, catalog number: BB0078)
20. D-glucose (BioShop, catalog number: GLU501)
21. D-sorbitol (BioShop, catalog number: SOR508)
22. DNA gel loading dye (New England Biolabs, catalog number: B7024S)
23. DpnI restriction enzyme (New England Biolabs, catalog number: R0176)
24. Ethylenediaminetetraacetic acid disodium salt, dihydrate (EDTA) (Bio Basic, catalog number: EB0185)
25. Ethanol, 95% purity (Greenfield Global, catalog number: P016EA95)
26. Ethanol, anhydrous (Greenfield Global, catalog number: P006EAAN)
27. Ethidium bromide (BioShop, catalog number: ETB444)
28. Glycerol (Bio Basic, catalog number: GB0232)
29. Iron chloride, hexahydrate (Bio Basic, catalog number: FD0201)
30. Isopropanol, min. 99.5% purity (Bioshop, catalog number: SO920)
31. L-arabinose (BioShop, catalog number: ARB222)
32. Lysozyme, egg white (BioShop, catalog number: LYS702)
33. Magnesium chloride, hexahydrate (BioShop, catalog number: MAG510)
34. Manganese (II) chloride, tetrahydrate (Sigma-Aldrich, catalog number: 805930)
35. Molybdic acid, sodium salt (Bio Basic, catalog number: MB0358)
36. Nickel (II) sulfate, hexahydrate (BioShop, catalog number: NIC700)
37. Phenol:chloroform:isoamyl alcohol, 25:24:1 (Fisher Scientific, catalog number: 15593031)
38. Polyethylene glycol 8000 (Fisher Scientific, catalog number: BP233-1)
39. Potassium acetate (Bio Basic, catalog number: PRB0438)
40. Potassium bromide (BioShop, catalog number: POB333)
41. Potassium chloride (Bio Basic, catalog number: PB0440)
42. Potassium chromate (Thermo Scientific Chemicals, catalog number: AC447201000)
43. Proteinase K solution (BioShop, catalog number: PRK222)
44. RNase A (QIAGEN, catalog number: 19101)
45. Selenious acid (Thermo Scientific Chemicals, catalog number: 211176)
46. Sodium acetate (Bio Basic, catalog number: SB1611)
47. Sodium bicarbonate (Bio Basic, catalog number: SB0482)
48. Sodium chloride (BioShop, catalog number: SOD004)
49. Sodium dodecyl sulfate (BioShop, catalog number: SDS001)
50. Sodium hydroxide (BioShop, catalog number: SHY700)
51. Sodium fluoride (BioShop, catalog number: SFL001)
52. Sodium nitrate (Bio Basic, catalog number: SD0484)
53. Sodium orthovanadate (BioShop, catalog number: SOV850)
54. Sodium phosphate, dibasic (BioShop, catalog number: SPD307)
55. Sodium phosphate, monobasic (BioShop, catalog number: SPM400)
56. Sodium sulfate (BioShop, catalog number: SOS513)
57. Thiamine hydrochloride, i.e., Vitamin B1 (Sigma-Aldrich, catalog number: T4625)
58. Tris hydrochloride (Bio Basic, catalog number: TB0103)
59. Yeast extract (BioShop, catalog number: YEX555)
60. Zinc sulfate, heptahydrate (Bio Basic, catalog number: ZB2906)
61. Zymolyase, 20,000 units/g (BioShop, catalog number: ZYM001)
Solutions
1. Biotin stock solution, 0.1% (w/v)
2. Boric acid stock solution, 0.1% (w/v)
3. Chilled isopropanol, 100% (v/v), -20 °C
4. Chilled ethanol, 70% (v/v), -20 °C
5. Cobalt (II) sulfate stock solution, 1% (w/v)
6. Copper (II) sulfate stock solution, 0.98% (w/v)
7. Cyanocobalamin stock solution, 0.1% (w/v)
8. D-sorbitol, 1 M
9. Ethylenediaminetetraacetic acid solution, 0.5 M, pH 8.0 (EDTA)
10. Glucose, 1 M
11. Glycerol, 50% (v/v)
12. Magnesium chloride, 1 M
13. Manganese (II) chloride stock solution, 18% (w/v)
14. Molybdic acid stock solution, 0.63% (w/v)
15. Nickel (II) sulfate stock solution, 0.27% (w/v)
16. Potassium chloride, 250 mM
17. Potassium chromate stock solution, 0.194% (w/v)
18. Selenious acid stock solution, 0.13% (w/v)
19. Sodium acetate, 3 M, pH 5.2
20. Sodium fluoride stock solution, 0.1% (w/v)
21. Sodium hydroxide solution, 1 M
22. Sodium orthovanadate stock solution, 0.184% (w/v)
23. Synthetic media lacking -Trp
24. Synthetic media lacking -His/Ura
25. Tris hydrochloride, 1 M, pH 8.0 (Tris-HCl)
26. Zinc sulfate stock solution, 2.2% (w/v)
DNA extraction
1. Buffer P1 (see Recipes)
2. Buffer P2 (see Recipes)
3. Buffer P3 (see Recipes)
4. CTAB lysis buffer (see Recipes)
Culturing E. coli
1. LB broth (see Recipes)
2. SOB broth (see Recipes)
3. SOC broth (see Recipes)
Culturing yeast
1. 2× YPAD broth (see Recipes)
2. Bacto-agar, 2% (w/v) (see Recipes)
3. Complete media (CM) glucose broth minus histidine and uracil (see Recipes)
Yeast assembly
1. SPEM solution (see Recipes)
2. Zymolase solution (see Recipes)
3. STC solution (see Recipes)
4. PEG-8000 solution (see Recipes)
5. SOS media (see Recipes)
Culturing P. tricornutum
1. NP stock, 500× (see Recipes)
2. L1 trace metals stock, 1,000× (see Recipes)
3. F/2 vitamin stock solution, 2,000× (see Recipes)
4. Anhydrous salts solution, 2× (see Recipes)
5. Hydrous salts solution, 2× (see Recipes)
6. L1 media (see Recipes)
Recipes
A. DNA extraction
1. Buffer P1
Store at 4 °C for < 12 months. Recipe made publicly available by QIAGEN.
Reagent Final concentration Amount
Tris-HCl (1 M, pH 8.0) 5.0 × 10-2 M 5 mL
EDTA (0.5 M, pH 8.0) 1.0 × 10-2 M 2 mL
RNAse A (100 mg/mL) 100 μg/mL 100 μL
ddH2O n/a 92.9 mL
Total n/a 100 mL
2. Buffer P2
Store at room temperature for < 24 months. Recipe made publicly available by QIAGEN.
Reagent Final concentration Amount
Sodium hydroxide 2.0 × 10-1 M 4 g
Sodium dodecyl sulfate 1% (w/v) 5 g
ddH2O n/a Top up to 500 mL
Total n/a 500 mL
3. Buffer P3
Store at room temperature for < 24 months. Recipe made publicly available by QIAGEN.
Reagent Final concentration Amount
Potassium acetate 3 M 147.2 g
ddH2O n/a Top up to 500 mL
Total n/a 500 mL
4. CTAB lysis buffer
Store at 4 °C for < 12 months. Recipe was obtained from Giguere et al. [9].
Reagent Final concentration Amount
Sodium chloride 1.4 M 4.1 g
Tris-HCl (1 M, pH 8.0) 2.0 × 10-1 M 10 mL
EDTA (0.5 M, pH 8.0) 5.0 × 10-2 M 5 mL
CTAB 2% (w/v) 1 g
RNAse A (100 mg/mL) 250 μg/mL 125 μL
ddH2O n/a Top up to 50 mL
Total n/a 50 mL
B. Culturing E. coli
1. LB broth
Sterilize by autoclaving, then store at room temperature for < 12 months. To make solid media, add 1.5 g of agar A for every 100 mL of LB (1.5%, w/v) prior to autoclaving.
Reagent Final concentration Amount
Bio-tryptone n/a 10 g
Sodium chloride 1.71 × 10-1 M 10 g
Yeast extract n/a 5 g
dH2O n/a Top up to 1,000 mL
Total n/a 1,000 mL
2. SOB broth
Mix all the components, excluding the magnesium chloride, then adjust the pH to 7.0 using NaOH and/or HCl as necessary. Sterilize by autoclaving and allow to cool to room temperature. Then, aseptically add 10 mL of sterile magnesium chloride solution. The magnesium chloride solution can be sterilized by autoclaving as well. SOB broth can be stored at room temperature for < 12 months.
Reagent Final concentration Amount
Bio-tryptone n/a 20 g
Yeast extract n/a 5 g
Sodium chloride 1.0 × 10-2 M 0.5 g
Potassium chloride, 250 mM 2.5 × 10-3 M 10 mL
dH2O n/a Top up to 990 mL*
Magnesium chloride, 1 M 1.0 × 10-3 M 10 mL
Total n/a 1,000 mL
3. SOC broth
Aseptically add 20% (w/v) filter-sterilized glucose to the sterile SOB broth. We recommend preparing 50 mL sterile aliquots of this media as it is very susceptible to contamination.
Reagent Final concentration Amount
SOB broth n/a 1,000 mL
Glucose, 1 M 2.0 × 10-2 M 20 mL
Total n/a 1,020 mL
C. Culturing yeast
1. 2× YPAD broth
Filter sterilize into a sterile glass bottle, then store at room temperature for < 12 months. D-glucose (i.e., dextrose) can burn if autoclaved, which will impact the growth of yeast. To make solid media, combine 2× YPAD with melted 2% (w/v) bacto-agar. Ensure the YPAD and bacto-agar have been equilibrated to 60 °C before combining in a 1:1 ratio to make 1% bacto-agar 1× YPAD plates. If using pre-mixed YPD agar (e.g., Sigma-Aldrich, catalog number: Y1500), combine 130 g/L of powder with dH2O and autoclave for 15 min at 121 °C to sterilize.
Reagent Final concentration Amount
Yeast extract n/a 20 g
Bacteriological peptone n/a 40 g
D-glucose 2.2 × 10-1 M 40 g
Adenine hemisulfate n/a 160 mg
dH2O n/a Top up to 1,000 mL
Total n/a 1,000 mL
2. Bacto-agar, 2% (w/v)
Autoclave to sterilize, ensuring that the glass bottle is no more than 80% full to avoid boiling over. Store at room temperature for < 24 months. The recipe below lists the volume we would prepare in a 500 mL glass bottle.
Reagent Final concentration Amount
Bacto-agar 2% (w/v) 8 g
dH2O n/a Top up to 400 mL
Total n/a 400 mL
3. Complete media (CM) glucose broth minus histidine and uracil
Adjust pH to 6.0, then autoclave to sterilize; store at room temperature for < 12 months. Only include D-sorbitol if plating spheroplasted yeast (i.e., during yeast assembly); omit when plating yeast with intact cell walls. To make solid media, add 2 g of Bacto-agar for every 100 mL of media (2%, w/v). Recipe is based on the manufacturer’s guidelines (Teknova) and Karas et al. [10].
Reagent Final concentration Amount
CM glucose broth minus histidine and uracil n/a 28.4 g
D-sorbitol 1.0 M 182 g
Adenine hemisulfate 8.69 × 10-4 M 160 mg
dH2O n/a Top up to 1,000 mL
Total n/a 1,000 mL
D. Yeast assembly
1. SPEM solution
Filter sterilize into a sterile glass bottle, then store at room temperature for < 12 months. Recipe was obtained from Karas et al. [10].
Reagent Final concentration Amount
D-sorbitol 1.0 M 182 g
EDTA (0.5 M, pH 8.0) 1.0 × 10-2 M 20 mL
Sodium phosphate, dibasic 1.47 × 10-2 M 2.08 g
Sodium phosphate, monobasic 2.32 × 10-3 M 0.32 g
dH2O n/a Top up to 1,000 mL
Total n/a 1,000 mL
2. Zymolyase solution
Sterilize using a syringe filter, then store in sterile 1.5 mL tubes at -20 °C for < 12 months. The efficiency of zymolyase noticeably drops with every freeze-thaw event, so we suggest storing aliquots of 45–90 μL, which is enough for 1–2 assembly reactions. Recipe was obtained from Karas et al. [10].
Reagent Final concentration Amount
Zymolyase 400 units/mL 200 mg
Tris-HCl (1 M, pH 8.0) 1.0 × 10-1 M 1 mL
Glycerol (50% v/v) 25% (v/v) 10 mL
ddH2O n/a 9 mL
Total n/a 20 mL
3. STC solution
Filter sterilize into a sterile glass bottle, then store at room temperature for < 12 months. It is optimal to store this solution as 15 mL aliquots in sterile conical tubes. Recipe was obtained from Karas et al. [10].
Reagent Final concentration Amount
D-sorbitol 1.0 M 182 g
Tris-HCl (1 M, pH 8.0) 1.0 × 10-2 M 10 mL
Calcium chloride, dihydrate 1.0 × 10-2 M 1.47 g
Magnesium chloride (1 M) 2.5 × 10-3 M 2.5 mL
dH2O n/a Top up to 1,000 mL
Total n/a 1,000 mL
4. PEG-8000 solution
Adjust pH to 8.0 with sodium hydroxide solution, then filter sterilize into a sterile glass bottle; store at 4 °C for < 12 months. PEG-8000 will depolymerize over time, thereby becoming less effective for yeast assembly. This process is exacerbated if the solution is left at room temperature. It is important to check the pH of this solution ahead of its use in assembly; when stored correctly, the pH should remain at or near 8.0 for up to a year. If the pH drops below this, dispose of the solution and remake it. Recipe was obtained from Karas et al. [10].
Reagent Final concentration Amount
PEG-8000 20% (w/v) 20 g
Calcium chloride, dihydrate 1.0 × 10-2 M 0.147 g
Magnesium chloride (1 M) 2.5 × 10-3 M 250 μL
Tris-HCl (1 M, pH 8.0) 1.0 × 10-2 M 1 mL
dH2O n/a Top up to 100 mL
Total n/a 100 mL
5. SOS media
Filter sterilize into a sterile glass bottle, then store at room temperature for < 12 months. It is optimal to store this solution as 15 mL aliquots in sterile conical tubes. Recipe was obtained from Karas et al. [10].
Reagent Final concentration Amount
D-sorbitol 1.0 M 182 g
Bacteriological peptone n/a 5 g
Yeast extract n/a 2.5 g
Calcium chloride, dihydrate 6.04 × 10-3 M 0.888 g
dH2O n/a Top up to 1,000 mL
Total n/a 1,000 mL
E. Culturing P. tricornutum
1. NP stock, 500×
Filter sterilize into a sterile glass bottle, then store at room temperature for < 24 months. Recipe was obtained from Karas et al. [11].
Reagent Final concentration Amount
Sodium nitrate 4.4 M 37.5 g
Sodium phosphate, monobasic 1.80 × 10-1 M 2.5 g
ddH2O n/a Top up to 100 mL
Total n/a 100 mL
2. L1 trace metals stock, 1,000×
Filter sterilize into a sterile glass bottle, then store at 4 °C for < 24 months. Recipe was obtained from Karas et al. [11].
Reagent Final concentration Amount
Iron chloride, hexahydrate 1.17 × 10-2 M 3.15 g
EDTA disodium salt, dihydrate 1.17 × 10-2 M 4.36 g
Copper (II) sulfate stock solution, 0.98% (w/v) 9.81 × 10-6 M 0.25 mL
Molybdic acid stock solution, 0.63% (w/v) 7.81 × 10-5 M 3.0 mL
Zinc sulfate stock solution, 2.2% (w/v) 7.65 × 10-5 M 1.0 mL
Cobalt (II) sulfate stock solution 1% (w/v) 3.56 × 10-5 M 1.0 mL
Manganese (II) chloride stock solution, 18% (w/v) 9.10 × 10-4 M 1.0 mL
Selenious acid stock solution, 0.13% (w/v) 1.01 × 10-5 M 1.0 mL
Nickel (II) sulfate stock solution, 0.27% (w/v) 1.03 × 10-5 M 1.0 mL
Sodium orthovanadate stock solution, 0.184% (w/v) 1.00 × 10-5 M 1.0 mL
Potassium chromate stock solution, 0.194% (w/v) 9.99 × 10-6 M 1.0 mL
ddH2O n/a Top up to 1,000 mL
Total n/a 1,000 mL
3. F/2 vitamin stock solution, 2,000×
Filter sterilize into a sterile glass bottle, then store at 4 °C for < 24 months. Recipe was obtained from Karas et al. [11].
Reagent Final concentration Amount
Thiamine hydrochloride 5.93 × 10-4 M 200 mg
Biotin stock solution, 0.1% (w/v) 4.09 × 10-5 M 10 mL
Cyanocobalamin stock solution, 0.1% (w/v) 7.38 × 10-7 M 1 mL
ddH2O n/a Top up to 1,000 mL
Total n/a 1,000 mL
4. Anhydrous salts solution, 2×
We use this immediately for making L1 media. If necessary, it can be filter sterilized and stored at 4 °C for < 12 months. Recipe was obtained from Karas et al. [11].
Reagent Final concentration Amount
Sodium chloride 8.38 × 10-1 M 24.5 g
Sodium sulfate 5.76 × 10-2 M 4.09 g
Potassium chloride 1.88 × 10-2 M 0.7 g
Sodium bicarbonate 4.76 × 10-3 M 0.2 g
Potassium bromide 1.68 × 10-3 M 0.1 g
Boric acid stock solution, 0.1% (w/v) 9.70 × 10-4 M 3 mL
Sodium fluoride stock solution, 0.1% (w/v) 1.43 × 10-4 M 300 μL
ddH2O n/a Top up to 500 mL
Total n/a 500 mL
5. Hydrous salts solution, 2×
We use this immediately for making L1 media. If necessary, it can be filter sterilized and stored at 4 °C for < 12 months. Recipe was obtained from Karas et al. [11].
Reagent Final concentration Amount
Magnesium chloride, hexahydrate 1.09 × 10-1 M 11.1 g
Calcium chloride, dihydrate 2.10 × 10-2 M 1.54 g
ddH2O n/a Top up to 500 mL
Total n/a 500 mL
6. L1 media
Adjust pH to 8.0 with sodium hydroxide solution, then filter sterilize into a sterile glass bottle; store at 4 °C for < 12 months. Note that this media lacks supplemental silica (sodium metasilicate nonahydrate) as it is not necessary for P. tricornutum growth. Recipe was obtained from Karas et al. [11].
Reagent Final concentration Amount
Anhydrous salts solution, 2× 1× 500 mL
Hydrous salts solution, 2× 1× 500 mL
NP stock solution, 500× 1× 2 mL
L1 trace metals solution, 1,000× 1× 1 mL
F/2 vitamin solution, 2,000× 1× 0.5 mL
Total n/a ~1,000 mL
Laboratory supplies
1. 0.2 mL PCR 8-strip tubes (FroggaBio, catalog number: STF-A120-S)
2. 1.5 mL tubes (FroggaBio, catalog number: 1210-001)
3. 15 mL conical tubes (FroggaBio, catalog number: TB15-500)
4. 50 mL conical tubes (FroggaBio, catalog number: TB50-500)
5. Bottle-top filters (≥ 500 mL capacity) with 0.2 μm PES membrane (Thermo Scientific, catalog number: 09-741-07)
6. Disposable hemocytometer, Neubauer-improved chamber (Fisher Scientific, SKC, catalog number: 22-600-100)
7. Disposable plastic cuvettes (VWR, catalog number: 97000-586)
8. Electrocuvettes, 2 mm (Fisher Scientific, catalog number: FB102)
9. Erlenmeyer flasks of various sizes (e.g., 100 mL, 250 mL)
10. EZ-10 Spin Column Plasmid DNA Miniprep kit (Bio Basic, catalog number: BS614)
11. EZ-10 Spin Column PCR Products Purification Kit (Bio Basic, catalog number: BS664)
12. Large-construct kit (QIAGEN, catalog number: 12462)
13. Liquid nitrogen (available at the institution)
14. Multiplex PCR kit (QIAGEN, catalog number: 206143)
15. Petri dishes, 100 × 15 mm (VWR, catalog number: 25384-088)
16. Porcelain mortar and pestle (Fisher Scientific, catalog numbers: FB961A and FB961K)
17. PrimeSTAR GXL DNA Polymerase kit (Takara, catalog number: R050A)
18. Two-sided disposable polystyrene cuvettes, 1.5–3.0 mL volume (VWR, catalog number: 97000-586)
19. Various sizes of glass bottles that can be sterilized
20. Pipette tips: 2 μL, 200 μL, 1,000 μL
Equipment
1. Biosafety cabinet (NuAire, model: LabGard ES NU-540)
2. Centrifuge with 15–50 mL tube capacity (e.g., Eppendorf, model: 5810, catalog number: 05-413-332)
3. Cryogenic dewar for liquid nitrogen
4. DeNovix spectrophotometer (DeNovix, model: DS-11, catalog number: DS-11)
5. Gel documentation system (Bio-Rad, model: ChemiDoc Imaging System, catalog number: 12003153)
6. Gel electrophoresis systems (Fisher Scientific, models: Owl EasyCast B1, B2 and B3, discontinued)
7. Gel power system (Fisher Scientific, catalog number: FB300Q)
8. Meker burner (Flinn Scientific, catalog number: AP1021)
9. Microcentrifuge with 1.5 mL tube capacity (Eppendorf, model: 5415C, discontinued)
10. pH probe (Sartorius, model: pHBasic+, discontinued)
11. Pipettes: P2, P20, P100, and P1000
12. Room or chamber at 18 °C with cool white light (i.e., blue-shifted) set to an intensity of 50–65 PPFD
13. Stationary/shaking incubators that can be set to 37 °C and 30 °C (Benchmark Scientific, catalog number: H1001-M)
14. Stir plate (Benchmark Scientific, catalog number: H4000-HS)
15. T100 thermal cycler (Bio-Rad, catalog number: 1861096)
Software and datasets
1. Benchling; free use, web-based platform (https://www.benchling.com/)
2. Image Lab; free use, application from Bio-Rad (version 6.1.0 build 7, standard edition)
3. Primer3web; free use, web-based platform (https://primer3.ut.ee/)
4. Sequenced and annotated P. tricornutum chloroplast genome; GenBank accession number EF067920.1, created by Oudot-Le Secq et al. [12]
Procedure
A. Designing the primers for PCR amplification of the chloroplast genome
1. Upload the sequenced and annotated chloroplast genome into Benchling or any other DNA manipulation software (e.g., SnapGene, Geneious). The P. tricornutum chloroplast genome can be accessed through GenBank (GenBank accession number EF067920.1; Oudot-Le Secq et al. [12]).
2. Roughly divide the genome into fragments that are between 5 and 20 kb and overlap by 400 bp.
Note: This overlapping region will be used to generate primers in step A3; the actual length of overlaps between fragments will range from 150 to 300 bp (Figure 1A and B).
The design principles we followed for splitting the P. tricornutum chloroplast genome into overlapping fragments are described in detail in our original article. Some considerations for the design include:
a. The cloning vector for whole-genome assembly should insert into a non-coding region of the genome; the chloroplast fragment termini should not overlap with each other at this insertion site (Figure 1A–D).
b. It is ideal to split the chloroplast genome into as few fragments as possible to increase assembly efficiency. The exact design approach used will depend on the genome structure and content, as well as the polymerase used for amplification. We were able to reliably amplify fragments as large as 18 kb with Takara PrimeSTAR GXL polymerase.
c. Avoid splitting the genome in repetitive regions, as this can lead to complications during yeast assembly (see Troubleshooting 1).
Figure 1. Designing primers to PCR-amplify the chloroplast genome as overlapping fragments. A) Primers are positioned within a 400 bp region of the chloroplast genome using Primer3, which generates optimized ~20 bp primers. The reverse primer will be used to amplify the upstream fragment, whereas the forward primer will be used to amplify the downstream fragment. B) The chloroplast genome should be split into fragments that overlap by 150–300 bp at every junction (box 1) except for where the cloning vector will integrate (box 2). Here, the chloroplast genome will be amplified using primers that will enable the integration of the cloning vector, which is split into two pieces. C) The cloning vector is amplified using primers that will add homologous sequences to the chloroplast genome at its termini. It is also split into two fragments that overlap in the HIS3 marker to reduce the occurrence of false positives during assembly. Here, the forward and reverse primers positioned at the vector’s termini contain 50 bp of homologous sequence to the integration region in the chloroplast genome. D) The chloroplast fragments are amplified using primers that add 20- to 40 bp of homologous sequences to the cloning vector at their respective termini. This will enable the integration of the cloning vector into a specific region in the chloroplast genome.
3. Copy the 400 bp (5' to 3' orientation) sequence where the fragments overlap and paste this into Primer3web. In the Product Size Ranges section, input 150–300, then click Pick Primers. This will generate a forward primer and reverse primer; use the forward primer for the “downstream” fragment, and the reverse primer for the “upstream fragment.” This will enable the amplification of fragments that overlap by 150–300 bp (Figure 1A), which is a suitable size for yeast assembly. Repeat this for every overlap region.
Note: Yeast can assemble fragments that have overlaps as small as 20 bp [13]; however, we recommend using larger overlaps where possible.
Troubleshooting: If Primer3 cannot identify any suitable primers, try selecting a larger region (e.g., 1,000 bp) with the same Product Size Ranges input. Alternatively, the overlap region can be shifted elsewhere; some regions of the chloroplast genome are particularly AT-rich, making it difficult to find optimal primers.
a. As mentioned above, the chloroplast fragments should not overlap at the region where the cloning vector will insert. For our design, we chose a non-coding region between fragments 7 and 8 as the integration site. Here, we designed forward and reverse primers that contained ~30 bp of homology to the chloroplast genome, and 40- to 60 bp of homology to the cloning vector termini (Figure 1D, Table 1). We also amplified the cloning vector using the same premise, such that there was ≥80 bp of homology between the respective fragment termini and cloning vector (Figure 1C, Table 1).
Note: We originally amplified the chloroplast fragments without the additional homology sequences to the cloning vector, but we recommend adding these to improve assembly efficiency.
b. We chose the cloning vector pPt0521S_URA (GenBank, accession number: KP745602.1) for our design because it contains BAC/YAC elements and an oriV sequence (see General note 9). The BAC/YAC elements enable plasmid replication, stability, and selection in both yeast and E. coli.
c. We used primers to split the cloning vector in the yeast HIS3 open reading frame using the primer optimization technique described in step A3. Splitting the cloning vector in the yeast selective marker can help reduce the number of false-positive transformants following assembly.
Table 1. Primers used for adding homology to the termini of the cloning vector and chloroplast fragments. Location refers to the coordinates of the chloroplast fragments relative to the reference genome (GenBank, accession number: EF067920.1; Oudot-Le Secq et al. [12]). Base pairs that are complementary to the chloroplast genome are shown in lowercase, whereas base pairs that are complementary to the cloning vector are shown in uppercase.
Amplicon Location Primers (5′ to 3′)
From To
Chloroplast genome fragment 7
77,856 91,605 Forward: tggaatttagttgggttacgc
Reverse: AGGGTTATGCAGCGGAAGATaaaaattcgttaattatttactt
aatacgaacatttaatttaatttatcaaaagttaaat
Cloning vector
fragment 1 91,605 91,606 Forward: ttttgataaattaaattaaatgttcgtattaagtaaataattaacgaatttttATC
TTCCGCTGCATAACCCTGCTTCGG
Reverse: TTCAGTGGTGTGATGGTCGT
Cloning vector fragment 2 Forward: CAGTAGCAGAACAGGCCACA
Reverse: taaaaatttactgaaaaaaatcaaataaacttagagaaagagtaattcttAA ACCAAAGCGGAGTGACTGCAACTAATGA
Chloroplast genome fragment 8
91,606 103,519 Forward: AATTTAATTTTCATTAGTTGCAGTCACTCCGCT TTGGTTTaagaattactctttctctaagtttatttgatttttttcag
Reverse: ttatcaccggcaaaaccttc
B. Obtaining overlapping fragments from the algal chloroplast genome
1. Isolate high molecular-weight (HMW) DNA (described in Giguere et al. [9])
a. We believe this DNA extraction method can be used for other algal species; however, we have only tested it with P. tricornutum, so we will write it as such.
b. Place a mortar and pestle into a -80 °C freezer hours ahead of attempting DNA isolation to ensure that it is sufficiently chilled.
c. Transfer 5 mL of a dense liquid culture of P. tricornutum (i.e., ≥ 6 × 106 cells/mL) into 50 mL of L1 media in a 250 mL glass flask. Repeat this across four flasks and grow until a density of 2–4 × 106 cells/mL is reached (~1 week). Place liquid cultures in a chamber at 18 °C with cool white light (i.e., blue-shifted) set to a photosynthetic photon flux density (PPFD) of around 70 μmol/s/m2.
Note: P. tricornutum cell density was estimated by counting cells with a hemocytometer. The culture was diluted 100× with L1 media. Then, 10 μL was pipetted into the counting chamber of a Neubauer-improved hemocytometer.
d. Transfer the cultures to four 50 mL conical tubes and spin at 3,000 RCF for 10 min at 4 °C. Decant the supernatant, ensuring that the pellet is not disrupted.
Pause point: The pellets can be left on ice for up to 2 h or frozen at -80 °C at this stage if needed. Frozen pellets can be kept for up to a month. It is ideal to flash-freeze the pellets if possible.
e. Resuspend each pellet with 1 mL of ice-cold TE buffer, then combine the resuspended cells into one conical tube.
f. Fill the pre-cooled mortar with a small volume of liquid nitrogen and begin adding the resuspended cells to it dropwise using a P1000 pipette. Add liquid nitrogen as needed to ensure the cells remain frozen as small beads.
g. Use the pre-cooled pestle to grind the frozen beads into a fine-grit powder, adding liquid nitrogen as needed to keep the cells frozen.
Note: The purpose of this step is to break down the exterior diatom cell wall, which is otherwise relatively impervious. Be sure to grind the cells sufficiently.
h. Transfer the frozen powder into a 15 mL conical tube and add 2 mL of CTAB lysis buffer and 10 μL (i.e., 200 μg) of proteinase K solution (20 mg/mL). Mix slowly using end-over-end inversion, then incubate at 37 °C for 15 min.
i. Gently mix the tube using end-over-end inversion once more, then place into a 37 °C incubator for another 15 min.
Note: As the cells lyse, the suspension will become a lighter green color.
j. Pellet the cell resuspension by spinning at 6,000 RCF for 5 min. Transfer the lysate to a new 15 mL conical tube.
k. Add one volume of phenol:chloroform:isoamyl alcohol (25:24:1) and mix gently by end-over-end inversion.
Caution: Phenol and chloroform are hazardous volatile chemicals; perform this step and subsequent steps in a ventilated fume hood with the appropriate personal protection equipment (PPE) until otherwise mentioned.
l. Centrifuge the sample at 6,000 RCF for 5 min, then carefully transfer the aqueous phase (i.e., top layer) to a new 15 mL conical tube.
m. Add one volume of chloroform:isoamyl alcohol (24:1) and mix gently by end-over-end inversion.
n. Centrifuge the sample at 6,000 RCF for 5 min, then carefully transfer 450 μL of the aqueous phase to a 1.5 mL tube. Repeat this as many times as necessary to remove as much of the aqueous phase as possible without transferring any of the interphase layer. After this point, the samples should no longer contain any chloroform, so all further steps can be conducted at a lab bench.
o. To the 1.5 mL tube(s), add a tenth volume of sodium acetate solution (3 M, pH 5.2) and two volumes of ice-cold 100% ethanol. Mix gently using end-over-end inversion.
Pause point: The sample(s) can be left at -80 °C for 1 h or -20 °C overnight to increase the yield of DNA.
p. Centrifuge the sample(s) at 16,000 RCF for 5 min, then decant the supernatant. Invert decanted tubes on a paper towel and dry until all residual ethanol has evaporated.
Optional: Use a chilled (i.e., ≤ 4 °C) centrifuge to increase the yield of DNA.
q. Resuspend the pellet with 100 μL of sddH2O and store at -20 °C.
Optional: Measure the concentration and purity of the genomic DNA using a fluorometer and/or spectrophotometer.
2. PCR amplification of fragments for whole-genome assembly
a. For the chloroplast fragments: Dilute the HMW DNA ~100 times in double-distilled water (ddH2O) that has been previously sterilized by autoclaving to ensure no nucleases are active. The concentration of the diluted DNA should be between 0.1 and 1 ng/μL.
b. For the cloning vector: Dilute a suitable cloning vector ~100–1,000 times in ddH2O so that there is less than 1 ng/μL of template DNA.
Note: In our original article, we used the pCC1BAC-derived cloning vector pPt0521S_URA for the PCR-based approach (GenBank, accession number: KP745602.1). The plasmid pINTO_7/8 is a SapI-domesticated version of pPt0521S_URA that we generated for the pre-cloned approach outlined in our original article; it can be obtained from Addgene (ID: 206431).
c. Prepare the PCR master mix (ddH2O, buffer, dNTPs, template DNA) according to the Takara PrimeSTAR GXL manual. This particular DNA polymerase kit was chosen as it has proven to be reliable at amplifying fragments as large as 18 kb (see General note 1).
d. Validate whether successful amplification took place by loading 1–2 μL the PCR products on a 1% agarose gel.
e. If amplification was successful, purify the PCR products using the BioBasic EZ-10 Spin Column PCR Products Purification kit, following the manufacturer’s guidelines.
3. Preparation of equimolar DNA mixtures for assembly
a. There are many different ways to estimate/measure DNA concentration. The easiest and perhaps most reliable is to use an agarose gel (Figure 2). To do this, load 1 μL of each assembly component into a 1% agarose gel along with 1 μL of an appropriate DNA ladder (e.g., 1 kb plus ladder, NEB).
Figure 2. 1% agarose gel where 1 μL of the fragments for chloroplast genome assembly (PCR-based approach) were visualized. Fragments 1–8 correspond to amplified regions of the P. tricornutum chloroplast genome, whereas the cloning vector fragments correspond to the amplified cloning vector, pPt0521S_URA. The NEB 1 kb ladder plus was used in lane 1 and serves as a reference for estimating the amount of DNA present in the subsequent lanes. Estimated mass values were generated in Image Lab.
b. Run the gel until reasonable separation has occurred, then image it using a gel documentation system. Download the .scn file and upload it onto a device that has the Image Lab software installed.
c. Open the .scn file in Image Lab and click on the Lanes and Bands button in the Analysis Tool Box section. Manually add the correct number of lanes onto the gel image, adjusting the frame as necessary, then manually add the bands for the ladder and sample lanes.
d. Go back to the Analysis Tool Box section and click on Quantity Tools. At the top of the selection, click Absolute and change the units to nanogram, leaving the regression method as linear.
e. Click on the top band of the ladder. The application will ask for a quantity value, which will depend on the specifications of the particular ladder used. For the 1 kb plus ladder, the band at 10 kb corresponds to 42 ng, the band at 6 kb corresponds to 50 ng, and so forth. Add the values for at least three ladder bands that have different masses.
f. Now, when you click on the bands that correspond to your samples, Image Scan will give a predicted value in nanograms. After this window pops up, record the value in a separate document, then press cancel.
Note: The actual nanograms of DNA will differ from the predicted value. What matters most here is to estimate the amounts of each sample relative to one another. Relative intensity can be gauged by the eye, but using a tool like Image Scan is more accurate.
g. Input the fragment length and estimated mass into a spreadsheet and perform the series of calculations demonstrated in Table 2. For whole-genome assembly, we created 40 μL assembly mixes. The assembly mix volume can be increased or decreased, so long as there is a sufficient amount (≥ 50 ng) of every assembly fragment; it is better to use more DNA than less. In our original article, we mistakenly reported that we used 50–400 μg of each fragment, when in fact, we used 50–400 ng.
h. Pipette the relative amounts of DNA into a 1.5 mL tube. Store at -20 °C until assembly is performed.
Note: It is recommended that at least two separate mixtures are prepared in case one of the fragments was not added properly.
Table 2. Calculations to estimate equimolar amounts of DNA from the agarose gel shown in Figure 2. Calculations are demonstrated in the first row containing data (i.e., Fragment 1 row). The relative proportion is calculated by dividing the value of length/mass in a cell by the sum of all values in the length/mass column. Then, the relative proportion value is multiplied by the anticipated final volume for the assembly reaction mixture, which is 40 μL in this example. This generates a proportional amount of DNA to add to the assembly mixture for each respective element. In practice, it is best to add a minimum of 1 μL of each assembly component to the mixture as demonstrated in the adjusted volumes column; this is to avoid pipetting error associated with microvolumes. For assembly of the P. tricornutum chloroplast genome, we amplified the genome as eight fragments that were assembled with the cloning vector pPt0521S_URA, which was split into two fragments (cloning vector fragments 1 and 2).
Assembly fragments Estimated mass (ng) Length (kb) Length/mass Relative proportion Volume to add to assembly mix (μL) Adjusted volumes (μL)
Fragment 1 31 14.5
31/14.5
= 0.468
0.468/4.47
= 0.105
40 × 0.105
= 4.2
4.2
Fragment 2 21 15.3 0.729 0.163 6.5 6.5
Fragment 3 149 17.3 0.116 0.026 1.0 1.0
Fragment 4 73 14.6 0.200 0.045 1.8 1.8
Fragment 5 11 17.6 1.600 0.358 14.3 14.3
Fragment 6 48 15.7 0.327 0.073 2.9 2.9
Fragment 7 18 13.8 0.767 0.172 6.9 6.9
Cloning vector, fragment 1 106 6.5 0.061 0.014 0.5 1.0
Cloning vector, fragment 2 300 8 0.027 0.006 0.2 1.0
Fragment 8 68 11.9 0.175 0.039 1.6 1.6
Total N/A N/A 4.47 N/A 40.0 41.2
C. Assembling the whole genome in S. cerevisiae (i.e., yeast assembly)
Adapted from the protocol described in Karas et al. [10]. Perform all steps aseptically.
1. Streak out a glycerol stock of S. cerevisiae strain VL6-48 onto 1% (w/v) bacto-agar YPAD plates and incubate at 30 °C until single colonies form (typically two days).
Note: Once struck out, plates containing VL6-48 can be parafilmed and kept at 4 °C for months. It is good practice to use recently passaged yeast ahead of assembly. The colonies should be white in color; if colonies appear pink, there is likely insufficient adenine hemisulfate present in the YPAD media.
2. At the beginning of the day, inoculate a single VL6-48 colony into 20 mL of 2× YPAD in a sterile 100 mL flask. Grow at 30 °C with 225 rpm shaking.
Note: If contamination issues occur, try adding ampicillin (100 μg/mL) to the media.
3. At the end of the day, measure the optical density at 600 nm wavelength (OD600). Dilute the culture into 50 mL of 2× YPAD such that the culture will reach an OD600 of 2.5–3.0 at the desired harvest time on the following day. Grow at 30 °C with 225 rpm shaking.
Note: The doubling time of VL6-48 can vary between 1.5 and 2.5 h. It is good practice to set up at least two additional cultures in case the yeast grows faster or slower than anticipated. Also, it is important to use 2× YPAD for culturing the yeast at this stage as growth in richer media has been shown to increase assembly efficiency.
a. We measure OD600 using 1 mL polystyrene cuvettes in a spectrophotometer. For accurate measurements, the OD600 value should fall between 0.2 and 1.0. At this stage, the starter culture is often below an OD600 of 1.0, so we use 1 mL of undiluted culture for measuring. If the culture is anticipated to have a density >1.0, we dilute it 2–10× using 2× YPAD (e.g., 10× = 100 μL of culture + 900 μL 2× YPAD).
b. When preparing additional cultures, we recommend setting them up so that one is at least a doubling unit ahead (i.e., more dense) and the other is at least a doubling unit behind (i.e., less dense) than the culture that is predicted to reach the desired OD.
c. Every 50 mL culture can be used to perform 10 assembly reactions. If including a positive and negative control (see General note 3), this means that eight separate constructs can be assembled.
4. When the culture reaches an OD600 of 2.5–3.0, transfer it to a 50 mL conical tube and centrifuge the cells at 2,500 RCF for 5 min at 10 °C. Decant the supernatant.
5. Resuspend the pellet in 20 mL of ddH2O by vortexing, then add an additional 30 mL of ddH2O. Invert to mix, then centrifuge at 2,500 RCF for 5 min at 10 °C. Decant the supernatant.
6. Resuspend the pellet in 20 mL of 1M D-sorbitol by vortexing.
Pause point: The resuspended cell pellet can be kept at 4 °C for up to 16 h without impacting assembly efficiency.
7. Add an additional 30 mL of 1M D-sorbitol. Invert to mix, then centrifuge at 2,500 RCF for 5 min at 10 °C. Decant the supernatant.
a. While centrifugation is taking place, prepare at least five cuvettes. In the first cuvette, mix 500 μL of ddH2O with 500 μL of 1M D-sorbitol. This will serve as a blank. Into the other cuvettes, add 800 μL of 1M D-sorbitol (mixture A) or ddH2O (mixture B). These cuvettes will be used to estimate the rate of the spheroplasting.
8. Resuspend the pellet in 20 mL of SPEM solution by vortexing. Add 30 μL of 2-mercaptoethanol and mix thoroughly by vortexing once more, then add 40 μL of zymolyase solution. Mix gently by end-over-end inversion, then place at 30 °C shaking at 75 rpm.
Critical: Once the zymolyase is added, it is crucial to be very gentle with the yeast. The spheroplasted cells are very sensitive to physical force; do not vortex the cells or shake too vigorously!
Caution: β-mercaptoethanol is a hazardous volatile chemical. We recommend performing this step and all proceeding steps in a biosafety cabinet until otherwise mentioned. Ensure that correct PPE is worn.
9. After 10 min has passed, gently transfer 200 μL of spheroplasted cells into the cuvette containing 800 μL of mixture A. Repeat this for mixture B. Then, place a piece of parafilm over the openings of the cuvettes and invert 3–5 times to mix.
10. Measure the OD600 and calculate the ratio of mixture A/mixture B. If the ratio is between 1.8 and 2.0, proceed to the next step. If the ratio is lower than this, continue to incubate the cells, checking on the optical density every 5–10 min (or as necessary).
Critical: Do not over-spheroplast the cells.
a. While measuring the OD, keep the cells stationary and at room temperature to avoid over-spheroplasting.
b. When first attempting this protocol, we suggest measuring and recording the A/B ratio before placing the culture into the incubator (i.e., time-point zero). These values can serve as a useful reference for gauging the rate of spheroplasting.
c. Spheroplasted cells are more apt to burst when placed in ddH2O compared to sorbitol; thus, as the reaction occurs, the ratio of mixture A/ mixture B should increase. If the cells over-spheroplast, this ratio will be greater than 2.0.
d. If an A/B ratio of 1.8–2.0 is not reached after 30 min, add another 15–30 μL of zymolyase solution. Incubate the cells as before and measure the optical density every 5 min.
11. Add 30 mL of 1M D-sorbitol to the cell mixture, then gently invert to mix. Centrifuge at 1,000 RCF for 5 min at 10 °C. Gently decant the supernatant.
Note: The cell pellet is very fragile and may break; be careful during this step and all subsequent steps.
Caution: Collect the supernatant, which contains 2-mercaptoethanol, in an appropriate vessel for hazardous disposal. The cell pellet should only contain trace amounts of 2-mercaptoethanol, so all further steps can be conducted at a lab bench.
12. Add 10 mL of 1M D-sorbitol, then gently resuspend the pellet by passing it 5–10 times through a 10 mL pipette.
Note: A P-1000 pipette can be used in lieu of a serological pipette, though the latter is more gentle as it has a wider opening.
13. Add an additional 30 mL of 1M D-sorbitol and gently invert to mix, then centrifuge as in step C12. Decant the supernatant gently.
a. While centrifugation occurs, prepare aliquot(s) of the PEG-8000 solution and equilibrate to 37 °C (see General note 2). The cell mixture will be resuspended with 1 mL of PEG-8000 solution at a later step.
14. Resuspend the pellet with 2 mL of STC solution, then incubate at room temperature for 10–20 min. As the cells incubate, remove the DNA assembly mix from -20 °C and allow it to thaw at room temperature. Prepare additional 1.5 mL tubes for the positive and negative control (see General note 3).
15. For each assembly reaction, combine 200 μL of the spheroplasted yeast with the respective volume of pre-mixed DNA. Mix by gently flicking the tube, then incubate at room temperature for 5 min.
16. Add 1 mL of 37 °C PEG-8000 solution to each assembly mixture and mix gently by inverting the tube 6–10 times. Incubate at room temperature for 15–20 min.
17. Centrifuge at 1,500 RCF for 7 min at room temperature. Carefully remove the supernatant with a P-1000 pipette.
18. Resuspend the pellet in 800 μL SOS media and incubate at 30 °C for at least 30 min.
Note: The cells can be left to incubate for up to 1 h without impacting assembly efficiency.
a. While the assembly mixtures incubate, melt the drop-out media required for assembly; we used -HIS/URA drop-out media for assembling the P. tricornutum chloroplast genome, but this will vary depending on which auxotropic selection markers are present in the construct design. Drop-out media can be melted by microwaving on a low-power setting to avoid bubbling over and burning. Ensure that 1M D-sorbitol is present in the media, as the spheroplasted cells will burst if it is absent.
b. Pour 8 mL aliquots of molten media into sterile 15 mL conical tubes, then place in a water bath set to 50 °C. Then, pour 20 mL aliquots of drop-out media into sterile Petri dishes (100 mm diameter). There should be two conical tubes and two plates per assembly reaction, as well as a conical tube and plate for each of the controls.
19. For each assembly reaction, combine 100 μL of the spheroplasted cells/DNA mixture with 8 mL of equilibrated drop-out media, then invert 3–5 times. Pour the mixture onto a Petri dish containing 20 mL of cooled drop-out media. Repeat this using the remaining 700 μL of cells (see General note 4).
a. The assembly plates will consist of a 20 mL bottom layer of drop-out media and an 8 mL top layer of drop-out media mixed with transformed yeast cells. The bottom layer of agar helps to prevent the plate from desiccating during incubation.
b. The spheroplasted yeast cells are very delicate. The agar matrix provides structural support for the cells during the initial stages of cell wall recovery, which is believed to enhance transformation efficiency.
c. Alternatively, the 8 mL of drop-out media mixed with transformed cells can be poured into an empty plate. After 1 day of incubation at 30 °C, 8 mL of liquid drop-out media can be added to the plate, which is then parafilmed to prevent desiccation. Here, colonies will grow into the top liquid, forming a pool of yeast transformants. To obtain single yeast colonies, it will be necessary to plate dilutions of the top liquid on suitable drop-out media plates after sufficient growth has occurred (i.e., > 4 days).
20. Once sufficiently dry (approximately 5 min), place the plates into a bag and move to a 30 °C incubator. For whole-genome assembly, colonies should emerge after 4–5 days. If colonies do not appear, see Troubleshooting 1.
D. Isolating and screening assembled constructs from S. cerevisiae
1. Isolating DNA from yeast transformants via alkaline lysis
a. Repatch at least 10 single transformants onto drop-out plates and incubate at 30 °C (Figure 3A, see General notes 5 and 6).
Note: At this point, the yeast cells will have recovered their cell walls; use drop-out agar that does not have D-sorbitol added, as it will interfere with cell growth if present (see Recipes).
When repatching, pick a single colony using a sterile pipette tip and make a 1–2 cm patch on the agar plate. Repeat this step until all 10 colonies have been patched on the plate. It is not necessary to streak out the transformants to obtain single colonies.
b. Once sufficient growth has occurred (approximately 2 days), inoculate each transformant into 5 mL of liquid drop-out media in 15 mL conical tubes. Place at 30 °C with shaking at 225 rpm overnight.
When inoculating, touch a sterile pipette tip to the end of the patch from step D1a and drop it into the liquid media.
c. The next day, pellet the cells by centrifuging at 3,000 RCF for 5 min at room temperature. Decant the supernatant.
d. Resuspend the pellet with 240 μL of P1 buffer, 5 μL of 2-mercaptoethanol, and 5 μL of zymolyase solution. Thoroughly mix and then transfer to a 1.5 mL tube.
Caution: 2-mercaptoethanol is a hazardous volatile chemical. We recommend performing this step and all future steps in a fume hood until otherwise mentioned. Ensure that correct PPE is worn.
e. Add 250 μL of P2 buffer and gently invert 6–10 times to mix. Incubate for 2 min at room temperature.
f. Add 250 μL of P3 buffer and thoroughly mix by inversion. Centrifuge at the maximum speed (e.g., 16,000 RCF) for 10 min at room temperature.
g. Transfer the supernatant (approximately 750 μL) to a clean 1.5 mL tube, then add 750 μL of -20 °C 100% isopropanol. Invert to mix, then centrifuge as in step D1f.
Pause point: The sample(s) can be left at -80 °C for 1 h or -20 °C overnight to increase the yield of DNA. Using a chilled centrifuge can also increase DNA yield.
h. Decant the supernatant, then add 500 μL of -20 °C 70% ethanol. Invert to mix, then centrifuge at maximum speed for 5 min at room temperature.
Optional: Use a chilled centrifuge to increase DNA yield.
i. Decant the supernatant and allow the DNA pellet to dry thoroughly. At this point, all traces of 2-mercaptoethanol should be gone.
j. Resuspend the pellet with 30–50 μL of ddH2O, then store at -20 °C.
Optional: Measure the concentration and purity of the DNA using a fluorometer and/or spectrophotometer. Alkaline lysis should yield a final concentration greater than 100 ng/μL.
2. Screening DNA for the assembly of complete chloroplast genomes
a. Prepare the QIAGEN multiplex (MPX) master mix according to the manufacturer’s guidelines (Figure 3B, see General note 7).
Note: The primer pairs used to screen for assembly of the whole genome span 6 of the 8 overlapping junctions between the chloroplast fragments. The design of these primers is described in detail in our original manuscript.
b. Use 1 μL of 10× diluted yeast DNA as the template for the reactions. For a positive control, use 1 μL of 100× diluted genomic DNA from P. tricornutum. For the negative control, use 1 μL of ddH2O.
Note: Including both controls is necessary. Any amplification of the negative control suggests contamination or carry-over of DNA during pipetting, see Troubleshooting 2.
c. Perform 30 cycles of the MPX reaction. Increase to 35 cycles if necessary.
Note: Increasing the number of cycles heightens the risk of false positives.
d. Load 2 μL of amplified MPX DNA on a 2% agarose gel with an appropriate DNA ladder (Figure 3C).
Note: The expected sizes for the amplicons fall within 100–700 bp, so it is necessary to use a 2% agarose gel for proper separation. If amplification fails for the yeast transformant DNA, see Troubleshooting 3.
3. Alternative screening method for assembly of the complete chloroplast genomes
a. If it is too cumbersome to perform alkaline lysis for several yeast transformants, an alternative method is to perform a rapid heat lysis of the cells.
b. Follow steps D1 (a and b) to passage yeast transformants. Instead of transferring the cells to liquid drop-out media, passage onto a plate for a second time.
c. Once sufficient growth has occurred (approximately 2 days), touch a pipette tip to the passaged yeast colony, then place the pipette tip into 10 μL of TE buffer in a microcentrifuge tube. Let it sit for 2–5 min before disposing of the pipette tip.
d. Vortex the tubes to sufficiently mix the contents, then incubate at 95 °C for 10 min.
Note: This heat treatment will lyse the cells, causing the release of DNA into the TE buffer.
e. Centrifuge the tubes until the cell debris has pelleted, then use 1 μL of the supernatant as template in the MPX reaction (step D2b).
Note: We only recommend using this method as a preliminary screen for successful transformants. The template DNA will not be of high quality and may lead to false negatives.
Figure 3. Passaging and screening yeast transformants post-assembly. A) At least ten individual yeast colonies are picked from the assembly plate and struck onto a fresh plate of drop-out media. After sufficient growth (~2 days), colonies are transferred to liquid drop-out media and grown overnight. The next day, DNA isolation via alkaline lysis is performed. B) Yeast transformants are screened using a multiplex PCR assay that spans the junctions where chloroplast fragments recombine (highlighted in pink). C) A 2% agarose gel where 2 μL of the multiplex PCR reactions were visualized. Transformants that demonstrate the expected multiplex banding pattern (C2–C10) suggest that the whole chloroplast genome has been assembled. The positive control consists of genomic DNA from P. tricornutum, and the negative control has no DNA added.
E. Transforming DNA from S. cerevisiae to E. coli via electroporation
Note: Perform all steps aseptically.
1. Use DNA from yeast transformants that screened positively for the presence of the whole chloroplast genome (Figure 4A).
Figure 4. Screening E. coli transformants post-transformation with yeast-assembled DNA. A) DNA isolated from yeast transformants that screen positively is electroporated into E. coli for further analysis. B) A 2% agarose gel where 2 μL of the multiplex PCR reactions were visualized. All E. coli transformants demonstrate the expected multiplex banding pattern, suggesting that the whole chloroplast genome has been successfully transformed. The positive control consists of total DNA isolated from P. tricornutum and the negative control has no DNA added.
2. Place sterile 2 mm electrocuvettes on ice.
Note: It is important to keep everything cold, as this prevents DNases from degrading the DNA during transformation.
3. Thaw a tube of EPI300 electrocompetent cells (Lucigen) on ice for approximately 10 min (see General note 8).
4. Once thawed, mix 25–50 μL of cells with 1–2 μL of isolated yeast DNA. Flick to mix, then incubate on ice for 5 min.
Note: It is ideal to use 50 μL of cells; however, 25 μL will suffice if needed. Use 2 μL of yeast DNA if the concentration appears low.
a. It is good practice to include positive and negative controls when conducting electroporation. Use 1 μL of plasmid DNA that contains the same marker(s) as the assembled construct for the positive control. The negative control will consist of cells with no DNA added.
5. Transfer the mixture to a chilled 2 mm electrocuvette, ensuring that the mixture sits evenly across the bottom of the cuvette and that there are no air bubbles present.
6. Pulse in an electroporator set to 2.5 kV with a capacitance of 25 μF and resistance of 200 Ω.
7. Add 1 mL of SOC media to the cuvette, then pipette up and down 3–5 times to resuspend the cells as they will be stuck to the bottom of the electrocuvette. Transfer as much as possible to a 1.5 mL tube, then place the mixture at 37 °C with shaking at 225 rpm for 1 h.
Note: The electrocuvettes can be washed and reused several times if properly cared for.
a. It is challenging to remove the whole reaction volume from the cuvette after resuspension. We find that 800–900 μL of resuspended cells are typically transferred to the 1.5 mL tube.
8. Individually plate 100 and 700 μL of the cells across two separate 1.5% agar (w/v) LB plates supplemented with chloramphenicol (15 μg/mL). Once the plates are dried, transfer to 37 °C. Colonies should emerge within 24 h (see General note 4).
F. Isolating and screening assembled constructs from E. coli
1. Isolating DNA from E. coli transformants via alkaline lysis
a. Inoculate a single E. coli colony into 5 mL of LB supplemented with chloramphenicol (15 μg/mL). Grow overnight at 37 °C shaking at 225 rpm.
Note: Do this for at least three E. coli colonies per transformation.
b. The next day, inoculate 500 μL of saturated culture into 5 mL of LB supplemented with chloramphenicol (15 μg/mL) and L-arabinose (100 μg/mL). Grow for 5 h at 37 °C on a shaker set to 225 rpm, ensuring that the cultures are well aerated (see General note 9).
c. Centrifuge 1–5 mL of the culture (1.5 mL is usually enough) at maximum speed for 2 min. Decant the supernatant.
d. Resuspend the pellet with 250 μL of P1 buffer, ensuring that the cells are mixed thoroughly.
e. Add 250 μL of P2 buffer, then invert 6–10 times to mix. Incubate at room temperature for 10 min.
f. Add 250 μL of P3 buffer and thoroughly mix by inversion. Centrifuge at the maximum speed (e.g., 16,000 RCF) for 10 min at room temperature.
g. Transfer the supernatant (approximately 750 μL) to a clean 1.5 mL tube, then add 750 μL of -20 °C 100% isopropanol. Invert to mix, then centrifuge as in step F1f.
Pause point: The sample(s) can be left at -80 °C for 1 h or -20 °C overnight to increase the yield of DNA. Using a chilled centrifuge can also increase DNA yield.
h. Decant the supernatant, then add 500 μL of -20 °C 70% ethanol. Invert to mix, then centrifuge at maximum speed for 5 min at room temperature.
Optional: Use a chilled centrifuge to increase DNA yield.
i. Decant the supernatant and allow the DNA pellet to dry thoroughly.
j. Resuspend the pellet with 30–50 μL of ddH2O, then store at -20 °C.
Optional: Measure the concentration and purity of the DNA using a fluorometer and/or spectrophotometer.
2. Screening DNA for the assembly of complete chloroplast genomes
Follow the same approach for screening yeast transformants (step D2) but dilute the template DNA 100–1,000× as it will be more abundant (Figure 4B).
3. Alternative screening method for assembly of complete chloroplast genomes
Follow the same approach for screening yeast transformants (step D3).
4. Isolating E. coli DNA for sequencing
Whole-plasmid sequencing is the gold standard for confirming that an assembly is correct. The DNA prepared through the alkaline lysis method works sufficiently for initial screens but is often too poor quality for sequencing due to the carryover of salts, RNA, and shorn genomic DNA during isolation. We used the QIAGEN large construct kit to purify the constructs from E. coli transformants that diagnostically appeared to have the correctly assembled genome.
Data analysis
The majority of data processing and analysis in our original article occurred post–yeast assembly. Here, we performed sequence alignments to investigate if any major rearrangements or mutations accrued when cloning the genome. We also assessed the burden and long-term stability of the chloroplast genome when maintained in E. coli under low-copy and high-copy number replication. For the methods presented in this article, there is very little data analysis to be conducted, as the focus is on performing a technically challenging method. Once the genome is assembled and diagnostically validated, we recommend performing next-generation sequencing of a few representative yeast and/or E. coli transformants to confirm that the entirety of the construct has been successfully cloned.
Validation of protocol
We repeated the assembly of the P. tricornutum chloroplast genome several times in our original article (Table 3). Though there were varying degrees of efficiency, in every iteration, there were at least 20 yeast colonies to screen post-assembly. We MPX-screened 10 yeast colonies per assembly method, with 90%–100% of the transformants demonstrating the expected banding pattern for assembly of the whole chloroplast genome. For each assembly method, DNA from two prospective yeast colonies was electroporated into E. coli. Three colonies were screened per transformation, with 100% of the transformants demonstrating the expected MPX banding pattern. We conducted whole-plasmid sequencing for at least one E. coli colony per assembly, all of which demonstrated the cloned chloroplast genome in its entirety. In the time since, we have used this method to assemble several permutations of the chloroplast genome, including halved-, quartered-, and two-third versions.
Table 3. Number of colony-forming units following assembly of the whole chloroplast genome in yeast. Derived from supplementary table S5 of Walker et al. [14].
Assembly approach Volume of spheroplasts plated
700 μL 100 μL
PCR-based approach 361 38
Pre-cloned approach, individual digestion of plasmids 47 2
Pre-cloned approach, one-pot digestion of plasmids 15 5
Combinatory approach for assembling pPt_Sap 20 2
General notes and troubleshooting
General notes
1. We followed these principles when amplifying fragments of the chloroplast genome using the Takara PrimeSTAR GXL kit:
a. When first testing primers, it is suitable to perform 10–25 μL reactions to conserve reagents.
b. When PCR-amplifying fragments greater than 5 kb, we use the fast protocol, which requires 2 μL of enzyme per 50 μL reaction. This amplifies DNA at a speed of 1 kb per 10 s.
c. If the PCR primers have been optimized using software like Primer3, we find that an annealing temperature of 60 °C works reliably. For primers that are not optimized, it may be necessary to vary the annealing temperature from as low as 50 °C, if no amplification occurs, and up to 68 °C if off-target bands are present.
d. For all of the PCR-amplified fragments, we began with 25 cycles and increased up to 35 cycles if amplification was insufficient. If you want to conserve reagents, you can place the reaction tube(s) back into the thermocycler for an additional 5–10 cycles. This only works if the tube(s) have not been frozen or left at room temperature for several hours.
2. PEG-8000 is a critical reagent in yeast assembly; it is important to purchase high-quality PEG from a reliable distributor. When testing out this protocol for the first time, we recommend purchasing PEG from more than one vendor to see which works best in your hands. As an anecdotal example, we observed nearly 10 times more yeast transformants when using the Fisher Scientific PEG (catalog number: BP233-1) compared to the Bio Basic PEG (catalog number: PB0433).
a. Make sure to store the PEG solution at 4 °C and test the pH of the solution ahead of use to ensure that it remained around a pH of 8.0 during storage.
b. When performing assembly, we recommend aseptically transferring an aliquot of PEG solution into a 15 mL conical tube that can then be equilibrated to 37 °C and disposed of after use. This is to avoid repetitively exposing the PEG solution to 37 °C as this can cause depolymerization over time. We also recommend keeping the PEG solution in an air-tight bottle as air exposure can lead to oxidation and depolymerization.
3. It is good practice to include positive and negative controls when performing assembly. Here, the negative control consists of spheroplasted yeast with no DNA added. The positive control consists of spheroplasted yeast mixed with at least 100 ng of a suitable control plasmid. This plasmid should contain the same selection marker(s) as the construct that you are assembling, as well as all other necessary elements for plasmid maintenance and replication in yeast and E. coli.
4. It can be beneficial to plate both 100 and 700 μL volumes of the cell mixtures. If assembly or transformation is very efficient, it will be easier for colonies to emerge when 100 μL of the reaction volume is plated as there will be less crowding. Conversely, if assembly or transformation is not efficient, then it is advantageous to have 700 μL of the reaction volume plated as this will give rise to more colonies. For the positive and negative controls, it is fine to plate the whole reaction volume (800 μL).
5. The cloned P. tricornutum chloroplast genome is ~130 kb, which is too large for most commercial spin-kit columns. We were able to successfully isolate the cloned genome from yeast and E. coli using alkaline lysis; however, this may not be possible for chloroplast genomes larger than this. For these cases, it may be necessary to use a specialized kit designed for the isolation of large plasmids.
6. Make sure to passage S. cerevisiae or E. coli transformants at least twice before conducting the multiplex screen. Any residual DNA left over from assembly or transformation can be carried over and, when screened, lead to false positive results. We recommend transferring as few cells as possible when passaging to avoid carry-over of this environmental DNA.
7. In a multiplex PCR reaction mixture, some primer pairs will amplify better than others. It may be necessary to increase or decrease the relative amounts of the primer pairs so that all of the amplicons have similar levels of amplification. This can be estimated by looking at the relative intensities of the amplicons after visualizing them on a 2% agarose gel.
a. An economical alternative to the QIAGEN MPX kit is the Takara SuperPlex kit. Though we did not use this in our original article, we have used the SuperPlex kit in the time since, and it appears to be as accurate and reliable as the QIAGEN kit.
b. It is important to include high-quality genomic DNA as a positive control for all multiplex reactions. It is also critical to include a negative control; here, we used 1 μL of TE buffer.
8. We performed transformation of the whole chloroplast genome using homemade electrocompetent EPI300 cells. Homemade cells are considerably less efficient than their commercial counterparts; with that being said, we were routinely able to get between 50 and 200 E. coli transformants, which was sufficient for our screening purposes. We mention this as commercial cells are notoriously expensive. We use the Warren [15] protocol to prepare large batches of electrocompetent EPI300 cells, which are stored in 50 μL aliquots at -80 °C until use.
9. We chose the EPI300 E. coli strain because it is highly competent and contains an arabinose-inducible mutant trfA gene integrated into its genome. When induced, trfA will lead to high-copy number expression of plasmids carrying an oriV (e.g., pCC1BAC-derived plasmids). This can be useful when trying to isolate large quantities of the construct from E. coli; however, high-copy level expression interferes with cell growth and can cause plasmid instability if sustained over several generations. Without supplemental arabinose, pCC1BAC plasmids are stably maintained as a single copy in EPI300. It is advisable to only induce an EPI300 culture for a few hours before harvesting the cells for DNA isolation. If your cloning vector does not contain an oriV, there is no need to supplement LB with L-arabinose.
Troubleshooting
1. Chloroplast genomes are not built the same across all photosynthetic eukaryotes; they can vary widely in size, gene content, repetitive regions, and more. If yeast assembly fails, here are a few things to consider:
a. Was there a positive control during assembly? The positive control provides a mechanism for ensuring that assembly worked; without this, it is hard to say if assembly failed due to the inability to correctly recombine the DNA fragments, over-spheroplasting, missing media elements (e.g., D-sorbitol), so on and so forth. When first attempting this method, we recommend preparing a large stock of a suitable plasmid that can be used as a positive control in every assembly. You should expect to see several hundreds to thousands of transformants on the positive control plate if the assembly was efficient. It is ideal to use a positive control plasmid that has the same yeast selection marker(s) as the construct you are trying to assemble.
b. What is the G+C content of the genome? It is known that DNA with low G+C content (≤30%) is challenging to clone and maintain in E. coli, whereas large fragments (≥100 kb) with high G+C content (≥40%) are challenging to clone in yeast [16]. If the G+C content is greater than 40%, it may be necessary to assemble portions of the genome at a time or add additional yeast replication origins throughout the genome [17]. Conversely, if the G+C content is lower than 30%, it may not be possible to transform and stably maintain the genome in E. coli.
c. Are there any toxic genes in the genome? Toxic genes encode proteins that serve a role in the host organism (or organelle, in this case) but cause disruption when expressed in other systems [18]. Also, regions with high A+T content can contain spurious open reading frames, which can be unexpectedly toxic when cloning [19]. This can be resolved by splitting the genome into multiple fragments and individually assembling each fragment with a suitable cloning vector in yeast [20]. The fragment containing the toxic gene should not give rise to many, if any, yeast and/or E. coli transformants compared to the other regions of the genome. The problematic fragment can be split further until the exact location of the toxic gene is identified, allowing for its targeted removal.
d. How big is the genome? The P. tricornutum chloroplast genome is small (117 kb) when compared to most plant and algal species. For genomes larger than this, it may be necessary to introduce additional origins of replication and other plasmid maintenance elements (e.g., markers) throughout the genome to ensure that it can be stably replicated and maintained in yeast and E. coli. This issue was encountered and overcome by O’Neill et al. [8] when cloning the 204 kb chloroplast genome of C. reinhardtii. A past study has shown that a 500 kb algal chromosome can be cloned in yeast and moved to E. coli, which is much larger than the vast majority of chloroplast genomes, so we believe this should not pose a major impasse [21].
e. How repetitive is the genome? The P. tricornutum chloroplast genome has two large inverted repeat (IR) regions that we had to account for when designing our assembly strategy, as these regions can easily recombine in yeast. Other chloroplast genomes may have larger IR regions or may contain multiple dispersive repetitive regions. It is important to take this into consideration when designing where the fragment termini are positioned and where the markers for yeast and E. coli will be integrated.
2. The QIAGEN MPX kit is very sensitive and can amplify trace amounts of DNA that are aerosolized during pipetting. If amplification occurs in the negative control, we suggest repeating the MPX reaction using a different set of pipettes. We also recommend using the best pipetting practices to prevent the aerosolization and carry-over of DNA; if possible, use barrier pipette tips (e.g., VWR, catalog number: 89082-364).
3. Alcohols, salts, and cell debris carried over during DNA isolation via alkaline lysis can interfere with the MPX DNA polymerase. If no amplification occurs when screening the yeast or E. coli transformants, try diluting the DNA another ten-fold and see if that restores amplification. If it does not resolve the issue, we recommend checking the DNA concentration and purity using a spectrophotometer. If the concentration and/or purity ratios are poor, try re-isolating the DNA and reperform the MPX reaction.
Acknowledgments
This work was supported by Natural Sciences and Engineering Research Council of Canada (RGPIN-2018-06172) awarded to B.J.K. The protocols described here, as well as the results, are derived from our original research paper [14].
Competing interests
The authors declare the following competing financial interest(s): B.J.K is Chief Executive Officer of Designer Microbes Inc. and holds Designer Microbes Inc. stock.
Ethical considerations
Not applicable for this protocol; no human and/or animal subjects were used.
References
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Article Information
Publication history
Received: Aug 30, 2024
Accepted: Nov 7, 2024
Available online: Nov 27, 2024
Published: Jan 20, 2025
Copyright
© 2025 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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Biological Engineering > Synthetic biology > Genetic modification
Molecular Biology > DNA > Chromosome engineering
Microbiology > Heterologous expression system > Saccharomyces cerevisiae
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Using Protein Painting Mass Spectrometry to Define Ligand Receptor Interaction Sites for Acetylcholine Binding Protein
AG Alexandru Graur
NE Natalie Erickson
NK Nadine Kabbani
Published: Vol 15, Iss 2, Jan 20, 2025
DOI: 10.21769/BioProtoc.5163 Views: 219
Reviewed by: Sébastien GillotinSarajo MohantaRohit Jain
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Original Research Article:
The authors used this protocol in ACS Chemical Neuroscience Jun 2024
Abstract
Nicotinic acetylcholine receptors (nAChRs) are a family of ligand-gated ion channels expressed in nervous and non-nervous system tissue important for memory, movement, and sensory processes. The pharmacological targeting of nAChRs, using small molecules or peptides, is a promising approach for the development of compounds for the treatment of various human diseases including inflammatory and neurogenerative disorders such as Alzheimer’s disease. Using the Aplysia californica acetylcholine binding protein (Ac-AChBP) as an established structural surrogate for human homopentameric α7 nAChRs, we describe an innovative protein painting mass spectrometry (MS) method that can be used to identify interaction sites for various ligands at the extracellular nAChR site. We describe how the use of small molecule dyes can be optimized to uncover contact sites for ligand–protein interactions based on MS detection. Protein painting MS has been recently shown to be an effective tool for the identification of residues within Ac-AChBP involved in the binding of know ligands such as α-bungarotoxin. This strategy can be used with computational structural modeling to identify binding regions involved in drug targeting at the nAChR.
Key features
• Identify binding ligands of nicotinic receptors based on similarity with the acetylcholine binding protein.
• Can be adapted to test various ligands and binding conditions.
• Mass spectrometry identification of specific amino acid residues that contribute to protein binding.
• Can be effectively coupled to structural modeling analysis.
Keywords: Ligand binding Nicotinic acetylcholine receptor Protein structure Mass spectrometry Drug discovery
Graphical overview
A summary of the workflow and main steps of the protein painting experimental design. Primary steps in the development and execution of the experiment. From left to right: The Ac-AChBP is produced in E. coli and then isolated using affinity chromatography; various dyes and enzymes are optimized prior to the protein paint experiment. Mass spectrometry analysis is used to define peptide fragments that are differentially produced in the painted experiment.
Background
Mammalian nicotinic acetylcholine receptors (nAChRs) are a family of ligand ion channels, which are widely expressed in the central and peripheral nervous systems. They play key roles in modulating neurotransmitter release, synaptic plasticity, and cognitive functions; they also regulate autonomic functions by mediating fast synaptic transmission in autonomic ganglia, impacting heart rate and blood pressure [1]. The pharmacological targeting of human nAChRs is a leading strategy for therapeutic drug development in the treatment of various nervous system disorders, including Alzheimer’s and neuropathic disease [2]. Homopentameric α7 nAChRs bind important proteins including pathogenic beta amyloids and various neurotoxins [3,4]. The structural similarity, solubility, and ligand-binding properties of the homopentameric invertebrate acetylcholine binding protein (AChBP) make it a suitable model for studying ligand binding at the extracellular domain of homopentameric nAChRs [5]. In this method paper, we describe the development of a protein painting mass spectrometry (MS) approach that can serve in the detection of protein binding domains within the AChBP.
Protein painting MS technology is a new biochemical strategy in the study of ligand binding to nAChRs, recently demonstrated by our study using AChBP derived from Aplysia californica (Ac-AChBP) [6,7]. Protein painting employs small molecular dyes that covalently “paint” the surface of multi-protein complexes and block access to enzymatic (e.g., trypsin) digestion during MS analysis (Figure 1) [8]. Through paint coverage, this method allows for the identification of ligand-binding regions useful for drug development at the nAChR site. In addition, protein painting MS complements other experimental tools, including computational structural modeling and site-directed mutagenesis of the nAChR ligand binding pocket [9]. The protein paint technique can be easily adapted and optimized (Figure 2) on a case-by-case basis, allowing for its utility in various studies. The components of the protein paint assay are widely available and include inexpensive protein dyes, common chemical reagents, and MS instrumentation that are accessible at research institutions. Results from protein painting can guide structure-activity relationship (SAR) studies in lead compound development during drug screening and may allow for the identification of additional allosteric binding sites within the nAChR. Lastly, protein painting MS can also assess for the potential binding of new ligands to the nAChR when there is no prior knowledge of interactions.
Figure 1. Identification of protein–protein interaction (PPI) sites through mass spectrometry (MS) protein painting. Protein painting utilizes the ability of various protein dyes to covalently bind to non-occupied (dye accessible) regions within a multi-protein complex. Dye inaccessible regions that correspond to possible PPI interaction sites are identified through MS analysis.
Figure 2. Flowchart showing the progression of experimental steps during the protein painting study
Materials and reagents
Biological materials
1. Escherichia coli BL21 (DE3) competent cells (Thermo Scientific, catalog number: EC0114)
Reagents
1. Invitrogen alpha-bungarotoxin conjugates (Thermo Fisher Scientific, catalog number: B1601)
2. Amyloid β-Protein 1-42 (Bachem, catalog number: 4014447)
3. 4-Nitrobenzenediazonium tetrafluoroborate (TCI, catalog number: N0137)
4. Disuccinimidyl suberate (DSS) (Thermo Fisher Scientific, catalog number: A39267)
5. Atto 425 NHS ester (Sigma-Aldrich, catalog number: 16805)
6. Dimethyl sulfoxide (DMSO) (Sigma-Aldrich, catalog number: D8418)
7. Anhydrous DMSO (Thermo Fisher Scientific, catalog number: D12345)
8. Choline (Acros Organics, catalog number: 110290500)
9. Nicotine (Sigma-Aldrich, catalog number: N3876)
10. Lysozyme (Thermo Fischer Scientific, catalog number: 89833)
11. Sequencing-grade trypsin (Promega, catalog number: V5111)
12. Sequencing-grade chymotrypsin (Promega, catalog number: V1061)
13. DNase (Sigma-Aldrich, catalog number: DN25-1G)
14. Complete EDTA-free mini protease inhibitor (Thermo Fisher Scientific, catalog number: A32955)
15. Protein ladder (Thermo Fischer Scientific, catalog number: 26619)
16. Phosphate-buffered saline (PBS) (VWR, catalog number: 45001-130)
17. Dithiothreitol (DTT) (Thermo Fischer Scientific, catalog number: R0861)
18. Urea (Thermo Fischer Scientific, catalog number: AC424581000)
19. Iodoacetamide (Sigma-Aldrich, catalog number: I1149)
20. Ammonium bicarbonate (Thermo Fischer Scientific, catalog number: A643)
21. Acetic acid (Thermo Fischer Scientific, catalog number: A38-500)
22. Acetonitrile (ACN) (Fischer Scientific, catalog number: A21-1)
23. Trifluoroacetic acid (TFA) (Thermo Fischer Scientific, catalog number: A116-1AMP)
24. LB broth (Thermo Fischer Scientific, catalog number: BP1426-2)
25. Ampicillin (AMP) (Thermo Fischer Scientific, catalog number: 11593027)
26. Isopropyl β-d-thiogalactopyranoside (IPTG) (Sigma-Aldrich, catalog number: I6758)
27. Tris base (Thermo Fischer Scientific, catalog number: BP152)
28. Sodium chloride (NaCl) (Thermo Fischer Scientific, catalog number: S25541)
29. Glycerol (Thermo Fischer Scientific, catalog number: J61059.AP)
30. Imidazole (Thermo Fischer Scientific, catalog number: 03196-500)
31. Triton X-100 (Sigma-Aldrich, catalog number: X100)
32. Coomassie Brilliant Blue G 250 (Sigma-Aldrich, catalog number: 1.15444)
33. Methanol (VWR, catalog number: BDH1135-4LG)
Solutions
1. Lysis buffer (see Recipes)
2. Resuspension buffer (Buffer A) (see Recipes)
3. Elution buffer (Buffer B) (see Recipes)
4. Final buffer (see Recipes)
5. Diazo dye (see Recipes)
6. NHS ester dye (see Recipes)
7. Sample buffer (see Recipes)
8. Coomassie solution (see Recipes)
Recipes
Notes:
1. All buffers should be autoclaved or filtered before use and kept at 4 °C.
2. All dyes should be prepared fresh, immediately before use and be protected from light exposure as much as possible.
1. Lysis buffer
20 mM Tris, pH 8.0
150 mM NaCl
10% glycerol
2. Resuspension buffer (Buffer A)
20 mM Tris, pH 8.0
150 mM NaCl
10% glycerol
10 mM imidazole
1% Triton X-100
3. Elution buffer (Buffer B)
20 mM Tris, pH 8.0
150 mM NaCl
10% glycerol
250 mM imidazole
1% Triton X-100
4. Final buffer
20 mM Tris, pH 8.0
150 mM NaCl
10% glycerol
5. Diazo dye
Solid 4-nitrobenzenediazonium tetrafluoroborate
1× PBS
Make 5 mg/mL stock. For each painted sample, you need 5 μL.
6. NHS ester dye
1 mg/mL Atto 425 NHS ester
Anhydrous DMSO
Make 5 mg/mL stock. For each painted sample, you need 5 μL.
7. Sample buffer
20% ACN
2% TFA
Sterile dd/diH2O
8. Coomassie solution
50% methanol
10% acetic acid
0.5% Coomassie Brilliant Blue G 250
Sterile dd/diH2O
Laboratory supplies
1. Eppendorf protein Lo-Bind 1.5 mL tubes (Eppendorf, catalog number: 022431081)
2. Sephadex G25 spin columns (Cytiva, catalog number: 27532501)
3. PierceTM C-18 spin columns (Thermo Fisher Scientific, catalog number: 89873)
4. Ni-charged IMAC column (Bio-Rad, catalog number: 12009287)
5. 30 kDa Amicon ultra centrifugal filter (Sigma-Aldrich, catalog number: UFC903008)
6. NuPAGE 4%–12% Bis-Tris gradient gel (Thermo Fisher Scientific, catalog number: NP0322BOX)
7. Graduated Erlenmeyer flasks (Sigma-Aldrich, catalog number: CLS4980250 and CLS49802L)
Equipment
1. Centrifuge 5810R (Eppendorf, catalog number: 022625501)
2. Sorvall WX+ Ultracentrifuge (Thermo Fisher Scientific, catalog number: 75000100)
3. Water bath (Thermo Fisher Scientific, catalog number: 15-474-18)
4. Sonicator (Thermo Fisher Scientific, catalog number: 15-338-281)
5. Tube rotator (Thermo Fisher Scientific, catalog number: 13-687-12Q)
6. Spectrophotometer (Cole Parmer, catalog number: EW-83059-10)
7. Exploris Orbitrap 480 coupled with an EASY-nLC 1200 HLPC system (Thermo Fisher Scientific, catalog number: BRE725533)
8. Reverse-phase PepMap RSLC C18 LC column (Thermo Fisher Scientific, catalog number: 164534)
9. Milli-Q IQ 7003/05/10/15 water purification system (Millipore Sigma, catalog number: C205110)
10. Autoclave
11. Incubated and refrigerated console shakers (Thermo Fisher Scientific, catalog number: SHKE435HP)
Software and datasets
1. Proteome Discoverer v2.3 (Thermo Fisher Scientific, 09/27/2019)
2. UCSF ChimeraX v1.6 (05/09/2023) (https://www.cgl.ucsf.edu/chimerax/)
3. Microsoft Excel 2016
4. RCSB Protein Data Bank (RCSB PDB) (https://www.rcsb.org/)
5. NCBI database, open access
6. Prism v10.2.0 (GraphPad, 03/26/2024)
Procedure
A. Make the Ac-AChBP
A1. Protein induction
1. Transform E. coli with recombinant Ac-AChBP with the N-terminal (His)6-tag in 200 mL of LB media with AMP (100 μg/mL) selection. Grow overnight at 37 °C with shaking.
2. Inoculate the above into 1.3 L of LB media at 37 °C. Save 2 mL (out of 200 mL volume) to use as a blank for the spectrophotometer reading.
3. Monitor cell growth by measuring the OD600 approximately 2 h after inoculation.
4. When the absorbance reaches 0.2, add IPTG to a final concentration of 1mM to induce protein expression. Incubate at 16 °C overnight.
A2. Cell harvest
1. Centrifuge at 6,000× g for 15 min at 4 °C.
2. Discard the supernatant, resuspend the pellet in fresh LB media, and repeat the centrifugation process.
3. Discard the supernatant and keep the cell pellet.
Optional: This pellet can be frozen at -80 °C for up to four months.
A3. Lysis and protein purification
1. Freshly prepare the lysis, resuspension, elution, and final buffers (see Recipes).
2. Resuspend the pellet in 50 mL of lysis buffer with 500 μL of 10 mg/mL DNase, 50 mg of lysozyme, and one Complete EDTA-free mini protease inhibitor tablet.
3. Sonicate the cells and then centrifuge at 39,000× g for 45 min. Collect the supernatant (lysis fraction).
4. Resuspend the pellet in 50 mL of resuspension buffer (Buffer A) with one Complete EDTA-free mini protease inhibitor tablet.
5. Sonicate the cells and then centrifuge at 39,000× g for 30 min. Collect the supernatant (load fraction) and the pellet (pellet fraction).
6. Equilibrate the Ni-NTA column with Buffer A.
7. Wash the column with 30–50 mL of Buffer A (supernatant from step A3.5) to remove nonspecific binding proteins. Save the flowthrough (wash fraction).
8. Elute the protein with 15–20 mL of Buffer B (elution fraction).
A4. Confirmation of protein expression
1. Perform a Bradford assay to determine protein concentration for each fraction (pellet, lysis, load, wash, and elution).
2. Load equal amounts of protein from each fraction onto a single SDS-PAGE gel and run the gel, ensuring complete separation of the molecular weight marker bands between 60 and 10 kDa.
3. Stain the gel using a Coomassie solution stain (see manufacturer’s instructions).
4. Confirm the expression of the Ac-AChBP subunit as the major protein product on the gel. The Ac-AChBP subunit is expected to run at ~28 kDa [6].
A5. Protein concentration and buffer exchange
1. Concentrate the eluted protein using a 30 kDa Amicon Ultra Centrifugal filter for 15 min at 4 °C.
2. Discard the flowthrough and refill the filter with the final buffer to remove imidazole.
3. Repeat the concentration process two times to complete the buffer exchange.
B. Protein paint
B1. Preparing the Ac-AChBP and ligand
1. Determine the appropriate number of samples and label a sterile Eppendorf protein Lo-Bind 1.5 mL tube for each. For example, use one tube for every individual Ac-AChBP unpainted condition, one tube for every Ac-AChBP painted condition, and one tube for every Ac-AChBP complex [Ac-AChBP and the tested ligand (α-Bungarotoxin)] painted condition.
Note: Additional ligands, including choline, nicotine, and amyloid β-Protein (1-42), can also be tested.
2. Prepare protein complexes in solution at 1:10 molar ratio of Ac-AChBP:ligand. For a 65 μL total solution, for example, prepare:
2.94 μL of Ac-AChBP
1.60 μL of α-Bungarotoxin
61.26 μL of 1× PBS
For control samples that do not contain a ligand, bring the final volume up to 65 μL with 1× PBS.
3. Place the protein paint complex on a rotator and incubate at room temperature for 1 h with gentle mixing at 20 rpm.
Note: Ensure that the solution is mixed well during the incubation.
4. Prepare the appropriate number of Sephadex columns as samples. Give each column a sharp downward shake to ensure that the resin moves to the bottom of the column. Remove the cap and place the column in a 2 mL Eppendorf collection tube.
5. Using a 20–200 μL pipette, remove any remaining Sephadex gel from inside of the cap and place it in the column.
6. Spin the Sephadex columns at 1,000× g for 1 min.
Note: Make sure there are no air pockets within the column as this might interfere with dye filtration.
7. Add 400 μL of MilliQ water and let sit at room temperature until the protein paint complex incubation step (step B1.3) is complete.
B2. Protein paint experiment
1. Add 5 μL of diazo dye (5 mg/mL, 1 mg in 200 μL of PBS, prepared fresh) to each painted protein sample. Add 5 μL of 1× PBS to each sample that will be unpainted.
2. Incubate samples for 30 min at room temperature on a rotator.
3. Spin all columns at 1,000× g for 1 min. Add 5 μL of 5 mg/mL NHS ester dye to the painted protein samples. Add 5 μL of DMSO to the unpainted sample.
Note: The volume must be consistent across the painted and unpainted groups during the entire experiment.
4. Place all samples on a tube rotator and incubate for 30 min at room temperature.
5. During the incubation period, add 400 μL of MilliQ water to each Sephadex column and centrifuge at 1,000× g for 2 min.
6. Remove the Sephadex columns from the 2 mL tubes and place in new labeled Eppendorf Lo-Bind 1.5 mL tubes. Ensure there are no air pockets or cracks in the resin before adding samples. If these are present, repeat the previous step by adding MilliQ water and spinning the columns once again at the same settings.
7. When the incubation is complete, transfer the entire 75 μL of sample from each tube to the appropriate Sephadex column.
Note: Carefully add the sample in the middle and avoid touching the resin with your pipette tip.
8. Centrifuge all samples at 1,000× g for 1 min. Throw away columns and retain the flowthrough.
B3. Denature proteins and add pre-digestion control
1. Add 0.8 μL of 250 μg/mL lysozyme stock to each sample.
2. Add 25 μL of 8 M urea to each sample, for a final volume of 100 μL and a final urea concentration of 2M.
3. Add 1.1 μL of 1 M DTT to each sample, for a final volume of 101.1 μL and a final DTT concentration of 10mM.
4. Incubate samples at 37 °C for 30 min using a water bath.
5. Add 12 μL of iodoacetamide to each sample, for a final volume of 113.1 μL and a final iodoacetamide concentration of 50 mM.
6. Incubate samples at room temperature in a dark place for at least 15 min.
B4. Trypsin digestion of protein complex
1. Add the following reagents according to the recipe below:
86.32 μL of MilliQ H2O (ultrapure water)
22 μL of ammonium bicarbonate
0.58 μL of trypsin*
*If your calculations for a 1:10 trypsin ratio result in a trypsin volume less than 0.5 μL, just add 0.5 μL.
This brings the final sample volume to 222 μL, final urea concentration to under 1M, ammonium bicarbonate to 50mM, and trypsin to 1:10 ratio.
2. Allow samples to digest overnight in a hot water bath at 37 °C.
3. Halt digestion by adding 5 μL of acetic acid, ~2% acetic acid in the final sample (or you can skip straight to preparing the C-18 columns).
4. Store samples at -20 °C until they can be prepped for mass spectrometry.
B5. Prepare C-18 columns
1. Thaw samples and add 75 μL of sample buffer to each sample.
2. Centrifuge samples at 1,000× g for 1 min. For technical replicates, divide each sample into two 150 μL aliquots in protein Lo-Bind 1.5 mL Eppendorf tubes. Alternatively, pass all 150 μL of sample through the column at one time.
3. Follow the C-18 spin column manufacturer’s directions to remove salt from samples.
4. For the final spin column step, elute each sample into a clean 1.5 mL tube.
5. Dry samples under nitrogen at 40 °C for roughly 10 min. Monitor closely to ensure no sample is blown out of the tube. Place dried peptides at -20 °C until mass spectrometry analysis can be performed.
Data analysis
1. LC/ESI MS: MS analysis was conducted in data-dependent mode with one full MS scan (60,000 resolving power) followed by MS/MS scans that target the top 20 abundant molecular ions fragmented by higher collisional dissociation (HCD). Tandem mass spectra were searched using Proteome Discover version 2.3 with SEQUEST against the NCBI E. coli databases and custom databases containing the recombinant protein sequence. Use a false discovery rate of 1% as the cutoff value for peptide spectrum matches (PSMs). In addition, label-free quantification via the use of the Minora algorithm within Proteome Discoverer can be applied to calculate protein abundances as described [10]. Peptides with less than two PSMs should be excluded from data analysis.
Note: Additional mass spectrometry parameters (e.g., methionine oxidation detection) are provided in Graur et al. [6].
2. Painted-to-unpainted abundance ratio measures and binding site validation:
Once data from at least three biological replicates is obtained, calculate separate abundance ratios for each identified peptide in the assay: the control ratio and the experimental ratio. For the control ratio, divide the peptide fragment's abundance reading in the painted Ac-AChBP sample by its abundance in the unpainted Ac-AChBP sample. For the experimental ratio, divide the peptide fragment's abundance reading in the painted Ac-AChBP + ligand sample by its abundance in the unpainted Ac-AChBP sample. If a ratio is less than 0.01, assign it a value of 0.01. Exclude peptide fragments with a control ratio greater than 0.25, which is likely due to dye inaccessibility. Perform a student’s t-test on the control and experimental ratios from all biological replicates to identify statistically significant peptide hits (p < 0.05).
Note: Peptide cleavage products of trypsin and chymotrypsin digestion can yield some peptide regions overlap. This is advantageous in mapping out various segments of the AChBP that may participate in ligand binding.
3. Visualization of the identified protein interaction sites:
According to the Protein Data Bank (PDB), the Ac-AChBP structure can be found in several ligand-bound states including the apo state (PDB ID:2BYN), in complex with epibatidine (PDB ID:2BYQ), or in complex α-bungarotoxin (PDB ID:7KOO). Protein structures can be visualized with software packages such as ChimeraX [11], and putative regions of ligand binding within the protein sequence can be identified from peptides obtained using the protein paint experiment. ChimeraX and other protein structural modeling tools are useful for visualizing regions involved in protein binding within individual subunits as well as the pentameric structure of the Ac-AChBP (Figure 3). This information can be used to determine the proximity of residues involved in ligand binding as well as the overall topology of the protein binding region identified in the protein paint MS assay.
Figure 3. Analysis of protein interaction sites within Ac-AChBP. A. The sequence of the Ac-AChBP (UniProt; Q8WSF8; 1–219) showing the location of structural residues (α helix, β sheet) as well as ligand binding sites (green). B–C. ChimeraX rendering of the Ac-AChBP pentamer (B) showing the location of ligand binding sites (green) within an individual subunit (blue) (C).
For a more detailed analysis, see Graur et al. [6].
Validation of protocol
This protocol has been used and validated across various experiments in the following research articles:
Graur et al. [6]. Protein Painting Mass Spectrometry in the Discovery of Interaction Sites within the Acetylcholine Binding Protein. ACS Chemical Neuroscience.
Haymond et al. [12]. Protein painting, an optimized MS-based technique, reveals functionally relevant interfaces of the PD-1/PD-L1 complex and the YAP2/ZO-1 complex. The Journal of Biological Chemistry.
Luchini et al. [8]. Protein painting reveals solvent-excluded drug targets hidden within native protein-protein interfaces. Nature Communications.
General notes and troubleshooting
Troubleshooting
Problem 1: Ac-AChBP protein production yield is too low for the protein paint experiment.
Possible cause: Low protein synthesis within BL21 cells.
Solution: Repeat the protein induction protocol and pool two (or more) post-induction cell lysate solutions prior to the affinity purification step. This should increase the total amount of Ac-AChBP available for the study.
Problem 2: Peptide ratio for painted-to-unpainted peptides is not significant.
Possible cause: Ligand concentration may not be optimal.
Solution: Calculate the appropriate ligand concentration that would saturate the binding site.
Problem 3: Peptide fragments identified by MS are too long or too short.
Possible cause: Digestion with enzymes such as trypsin may create tryptic peptide fragments of various lengths depending on the amino acid composition of the full-length protein. In some cases, peptide fragments may therefore be too long or too short for meaningful analysis within the protein paint study.
Solution: Prior to the protein paint experiment, in silico tools such as Peptide Cutter can be used to predict the size and location of enzymatic digestion and peptide fragments generated by peptidase cleavage with certain enzymes. In addition, in some experiments, it may be best to use more than one enzyme for protein digestion.
Acknowledgments
We thank Mr. Shahzaman Saeed for his assistance with the figures. This study was supported by an Alzheimer's and Related Diseases Research Award Fund (ARDRAF) grant to N.K. This protocol was originally published in Graur et al. [6].
Competing interests
The authors declare no competing interests.
References
Changeux, J. P. (2012). The Nicotinic Acetylcholine Receptor: The Founding Father of the Pentameric Ligand-gated Ion Channel Superfamily. J Biol Chem. 287(48): 40207–40215.
Bouzat, C., Lasala, M., Nielsen, B. E., Corradi, J. and Esandi, M. D. C. (2017). Molecular function of α7 nicotinic receptors as drug targets. J Physiol. 596(10): 1847–1861.
Bencherif, M., Lippiello, P. M., Lucas, R. and Marrero, M. B. (2011). Alpha7 nicotinic receptors as novel therapeutic targets for inflammation-based diseases. Cell Mol Life Sci. 68(6): 931–949.
Graur, A., Sinclair, P., Schneeweis, A. K., Pak, D. T. and Kabbani, N. (2023). The human acetylcholinesterase C-terminal T30 peptide activates neuronal growth through alpha 7 nicotinic acetylcholine receptors and the mTOR pathway. Sci Rep. 13(1): 11434.
Nemecz, Ã. and Taylor, P. (2011). Creating an α7 Nicotinic Acetylcholine Recognition Domain from the Acetylcholine-binding Protein. J Biol Chem. 286(49): 42555–42565.
Graur, A., Haymond, A., Lee, K. H., Viscarra, F., Russo, P., Luchini, A., Paige, M., Bermudez-Diaz, I. and Kabbani, N. (2024). Protein Painting Mass Spectrometry in the Discovery of Interaction Sites within the Acetylcholine Binding Protein. ACS Chem Neurosci. 15(11): 2322–2333.
Hansen, S. B., Sulzenbacher, G., Huxford, T., Marchot, P., Bourne, Y. and Taylor, P. (2006). Structural Characterization of Agonist and Antagonist-Bound Acetylcholine-Binding Protein From Aplysia californica. J Mol Neurosci. 30: 101–102.
Luchini, A., Espina, V. and Liotta, L. A. (2014). Protein painting reveals solvent-excluded drug targets hidden within native protein–protein interfaces. Nat Commun. 5(1): 4413.
Ho, T. N. T., Lee, H. S., Swaminathan, S., Goodwin, L., Rai, N., Ushay, B., Lewis, R. J., Rosengren, K. J. and Conibear, A. C. (2021). Posttranslational modifications of α-conotoxins: sulfotyrosine and C-terminal amidation stabilise structures and increase acetylcholine receptor binding. RSC Med Chem. 12(9): 1574–1584.
Horn, D. M., Uecker, T., Fritzemeier, K., Tham, K., Paschke, C., Berg, F., Pfaf, H., Jiang, X., Li, S. and Lopez-Ferrer, D. (2016). New Method for Label-Free Quantification in the Proteome Discoverer Framework. Poster. Thermo Fisher Scientific, San Jose, CA, USA; Thermo Fisher Scientific, Bremen, Germany.
Pettersen, E. F., Goddard, T. D., Huang, C. C., Meng, E. C., Couch, G. S., Croll, T. I., Morris, J. H. and Ferrin, T. E. (2021). UCSF ChimeraX: Structure visualization for researchers, educators, and developers. Protein Sci. 30(1): 70–82.
Haymond, A., Dey, D., Carter, R., Dailing, A., Nara, V., Nara, P., Venkatayogi, S., Paige, M., Liotta, L., Luchini, A., et al. (2019). Protein painting, an optimized MS-based technique, reveals functionally relevant interfaces of the PD-1/PD-L1 complex and the YAP2/ZO-1 complex. J Biol Chem. 294(29): 11180–11198.
Article Information
Publication history
Received: Aug 21, 2024
Accepted: Nov 20, 2024
Available online: Dec 5, 2024
Published: Jan 20, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Neuroscience > Cellular mechanisms > Receptor-ligand binding
Biochemistry > Protein > Interaction
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Mouse-derived Synaptosomes Trypsin Cleavage Assay to Characterize Synaptic Protein Sub-localization
JS Jasmeet Kaur Shergill
DT Domenico Azarnia Tehran
Published: Vol 15, Iss 2, Jan 20, 2025
DOI: 10.21769/BioProtoc.5164 Views: 238
Reviewed by: Marion HoggXiaoliang Zhao Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Science Advances May 2022
Abstract
Neurons communicate through neurotransmission at highly specialized junctions called synapses. Each neuron forms numerous synaptic connections, consisting of presynaptic and postsynaptic terminals. Upon the arrival of an action potential, neurotransmitters are released from the presynaptic site and diffuse across the synaptic cleft to bind specialized receptors at the postsynaptic terminal. This process is tightly regulated by several proteins at both presynaptic and postsynaptic sites. The localization, abundance, and function of these proteins are essential for productive neurotransmission and are often affected in neurological and neurodegenerative disorders. Here, we outline a method for purifying mouse synaptosomes and using limited tryptic digestion to assess the subcellular localization of synaptic proteins. During synaptosomes purification, presynaptic terminals reseal and are protected from proteolysis, while postsynaptic proteins remain susceptible to tryptic cleavage. These changes can easily be evaluated by western blot analysis. This approach offers a straightforward and reliable method to evaluate the subcellular localization of synaptic proteins based on their proteolytic sensitivity, providing valuable insights into synaptic physiology and pathology.
Key features
• Builds upon the method developed by Boyken et al. [1] and introduces the use of isolated mouse synaptosomes to assess synaptic protein sub-localization.
• Limited tryptic digestion differentiates between presynaptic and postsynaptic proteins based on proteolytic sensitivity.
• Requires standard biochemical reagents and western blotting equipment and can be completed in two/three days, including synaptosome purification and western blot analysis.
Keywords: Synaptosomes Trypsin Fractionation Synapse Presynaptic Postsynaptic Neurotransmission Western blotting
Graphical overview
Overview of the synaptosomes trypsin cleavage assay. This protocol describes the isolation of synaptosomes from mouse brain tissue, followed by limited trypsin digestion to assess the compartmental localization of synaptic proteins. Synaptosomes, which are isolated via differential centrifugation, consist of resealed presynaptic terminals that are protected from proteolysis, while the exposed postsynaptic compartments are accessible to trypsin and undergo proteolytic cleavage. Following digestion, samples are analyzed using SDS-PAGE and western blotting. Proteins of interest are probed using specific antibodies to determine whether they are presynaptic or postsynaptic. Presynaptic proteins (e.g., synaptophysin, SNAP-25) remain intact, while postsynaptic proteins (e.g., GluA1, GluN2A) are cleaved.
Background
Neurotransmission is the fundamental process by which neurons communicate and relay information, enabling neuronal circuits to function effectively, thus governing higher-order functions such as memory formation and storage [2]. This communication occurs at specialized structures called synapses, which consist of presynaptic and postsynaptic terminals [2–5]. In the presynaptic neuron, upon the arrival of an action potential, neurotransmitter-containing synaptic vesicles fuse with the presynaptic membrane, releasing their contents into the synaptic cleft [2–5]. These neurotransmitters diffuse across the cleft and bind to specialized receptors at the postsynaptic terminal, triggering the transmission of signals in the postsynaptic neuron [2–5]. This entire process is tightly regulated by a complex set of proteins [6–7]. Understanding the localization, abundance, and function of these proteins is essential for uncovering the mechanisms that regulate synaptic transmission, which can be altered in a variety of neurological and neurodegenerative disorders, such as Alzheimer's disease, Parkinson’s disease, and schizophrenia [8–9].
Several methodologies have been developed to study the protein composition of synapses [10]. Techniques such as immunohistochemistry and mass spectrometry have been widely used to map synaptic proteins [11–14]. Although these methods offer valuable insights into synaptic protein localization and abundance, they lack the resolution to distinguish between presynaptic and postsynaptic sites, which are separated by the synaptic cleft (approximately 20 nm wide). More recently, super-resolution microscopy has offered this level of resolution, but it often involves significant sample preparation time and is less suitable for high-throughput analysis [15–16]. To address some of these limitations and to isolate synaptic vesicle docking complexes, a method involving the use of synaptosomes combined with limited tryptic digestion has been developed by Boyken et al. [1]. This approach allows for the study of synaptic proteins in a more high-throughput and compartment-specific manner. Synaptosomes are partially resealed structures derived from isolated nerve terminals from mouse brains, in which presynaptic terminals remain sealed and intact while the postsynaptic terminal is left unprotected. By applying limited tryptic digestion to these synaptosomes, presynaptic proteins are protected from proteolysis, as they remain inaccessible to trypsin, while postsynaptic proteins are vulnerable to cleavage [1]. This method thus enables the distinction between presynaptic and postsynaptic proteins by western blot analysis, offering an efficient way to assess their localization.
To our knowledge, the protocol presented here has been used to study the molecular architecture of active zones [1], to confirm the postsynaptic localization of Synaptotagmin-3 [17], to assess the large postsynaptic pools of CALM [18] and intersectin1 (ITSN1) [19], and to confirm the postsynaptic localization of the total and phosphorylation forms of p85S6K over p70S6K, which instead appear to be mainly presynaptic [20]. Nevertheless, this protocol has the potential to be applied in various contexts in combination with mass spectrometry to enable a more precise understanding of the molecular components of synapses.
Materials and reagents
Biological materials
1. Mouse (Mus musculus, C57BL/6J) brain (preferably P60 or above)
Reagents
1. Sucrose (ROTH, catalog number: 9097.1)
2. HEPES (ROTH, catalog number: 6763.2)
3. Trypsin (Sigma-Aldrich, catalog number: T1005)
4. Bradford reagent (Sigma-Aldrich, catalog number: B6916)
5. Tris (ROTH, catalog number: AE15.6)
6. Glycerin (ROTH, catalog number: 3783.3)
7. SDS (ROTH, catalog number: 2326.2)
8. 2-Mercaptoethanol (Sigma Aldrich, catalog number: M3148)
9. Bromophenol blue (ROTH, catalog number: T116.1)
10. HCl (ROTH, catalog number: 4625.1)
11. Glycine (ROTH, catalog number: 3790.3)
12. Methanol (ROTH, catalog number: 8388.6)
13. Ponceau S (ROTH, catalog number: 5938.1)
14. Acetic acid (ROTH, catalog number: 3738.2)
15. Disodium hydrogen phosphate (Na2HPO4) (ROTH, catalog number: T876.1)
16. Potassium dihydrogen phosphate (KH2PO4) (ROTH, catalog number: 3904.1)
17. Sodium chloride (NaCl) (ROTH, catalog number: 3957.1)
18. Potassium chloride (KCl) (ROTH, catalog number: 6781.1)
19. Tween-20 (Polysorbat) (Fischer Scientific, catalog number: T/4206/60)
20. Intercept blocking buffer (LI-COR, catalog number: 927-60001)
21. ROTIPHORSE Gel 30 (37.5:1), ready-to-use acrylamide (ROTH, catalog number: 3029.1)
22. Ammonium peroxydisulfate (APS) (ROTH, catalog number: 9592.5)
23. TEMED (ROTH, catalog number: 2367.1)
24. Bovine serum albumin fraction V (BSA) (Sigma-Aldrich, catalog number: 1.12018.0100)
25. 2-propanol (isopropanol) (VWR, catalog number: 20842.323)
26. PageRuler prestained protein ladder (Thermo Scientific, catalog number: 26616)
27. Blotting paper (MACHEREY-NAGEL, catalog number: MN 827 B)
28. Nitrocellulose membrane (Amersham, catalog number: 10600004)
29. Anti-GluA1 antibody (Merk Millipore, catalog number: MAB2263)
30. Anti-GluA2 antibody (Merk Millipore, catalog number: MAB397)
31. Anti-GluN2A antibody (Cell signalling, catalog number: #4205)
32. Anti-GluN2B antibody (Cell signalling, catalog number: #4207)
33. Anti-Homer1 antibody (Synaptic Systems, catalog number: 160003)
34. Anti-Synaptotagmin1 antibody (SYT1) (Synaptic Systems, catalog number: 105103)
35. Anti-Synaptophysin1 antibody (SYP1) (Synaptic Systems, catalog number: 101011)
36. Anti-Snap25 antibody (Synaptic Systems, catalog number: 111011)
37. IRDye 800CW goat anti-Rabbit IgG secondary antibody (LI-COR, catalog number: 926-32211)
38. IRDye 680RD goat anti-Mouse IgG secondary antibody (LI-COR, catalog number: 926-68070)
Solutions
1. Sucrose buffer (see Recipes)
2. BSA solution for Bradford assay (see Recipes)
3. Trypsin solution (see Recipes)
4. 6× Loading sample buffer (LSB) (see Recipes)
5. 4× separating gel buffer (see Recipes)
6. 4× stacking gel buffer (see Recipes)
7. 10× running buffer (see Recipes)
8. 1× running buffer (see Recipes)
9. 10× transfer buffer (see Recipes)
10. 1× transfer buffer (see Recipes)
11. Ponceau S staining solution (see Recipes)
12. 10× PBS (see Recipes)
13. 1× PBS (see Recipes)
14. 1× PBST (see Recipes)
15. Blocking solution (see Recipes)
16. 10% APS solution (see Recipes)
Recipes
1. Sucrose buffer
Reagent Final concentration Quantity or Volume
Sucrose 0.32 M 10.95 g
HEPES 5 mM 0.119 g
Water n/a 100 mL
Total n/a 100 mL
Adjust the pH to 8.0 before reaching the final volume.
2. BSA stock solution for Bradford assay
Reagent Final concentration Quantity or Volume
BSA 2 mg/mL 2 mg
Sucrose buffer n/a 1 mL
Total n/a 1 mL
3. Trypsin solution
Reagent Final concentration Quantity or Volume
Trypsin 0.1 mg/mL 1 mg
Water n/a 10 mL
Total n/a 10 mL
Store the solution on ice after preparation.
4. 6× loading sample buffer (LSB)
Reagent Final concentration Quantity or Volume
Tris pH 6.8 375 mM 2.268 g
Glycerin 50% 25 mL
2-mercaptoethanol 10% 5 mL
SDS 10% 5 g
Bromophenol blue 0.03% 15 mg
Total n/a 50 mL
Heat the solution to 55 °C to dissolve the SDS, then add 2-mercaptoethanol at the end.
5. 4× Separating gel buffer
Reagent Final concentration Quantity or Volume
Tris pH 8.8 1.5 M 90.86 g
SDS 0.4% 2 g
Water n/a 500 mL
Total n/a 500 mL
Tris base pH level is adjusted to 8.8 using HCl. Add SDS at the end.
6. 4× Stacking gel buffer
Reagent Final concentration Quantity or Volume
Tris pH 6.8 0.5 M 30.29 g
SDS 0.4% 1 g
Water n/a 250 mL
Total n/a 250 mL
Tris base pH level is adjusted to 6.8 using HCl. Add SDS at the end.
7. 10× Running buffer
Reagent Final concentration Quantity or Volume
Tris pH 8.4 250 mM 30.29 g
Glycine 1.92 M 144.13 g
SDS 1% 10 g
Water n/a 1,000 mL
Total n/a 1,000 mL
8. 1× Running buffer
Reagent Final concentration Quantity or Volume
10× Running buffer 1× 100 mL
Water n/a 900 mL
Total n/a 1,000 mL
9. 10× Transfer buffer
Reagent Final concentration Quantity or Volume
Tris 250 mM 30.29 g
Glycine 1.92 M 144.13 g
Water n/a 1,000 mL
Total n/a 1,000 mL
10. 1× Transfer buffer
Reagent Final concentration Quantity or Volume
10× Transfer buffer 1× 100 mL
Methanol 20% 200 mL
Water n/a 700 mL
Total n/a 1,000 mL
11. Ponceau S staining solution
Reagent Final concentration Quantity or Volume
Ponceau S 0.5% 1.25 g
Acetic acid 1% 2.5 mL
Water n/a 250 mL
Total n/a 250 mL
12. 10× PBS
Reagent Final concentration Quantity or Volume
Na2HPO4 100 mM 11.49 g
KH2PO4 18 mM 2.45 g
NaCl 1.4 M 82.82 g
KCl 27 mM 2.01 g
Water n/a 1,000 mL
Total n/a 1,000 mL
13. 1× PBS
Reagent Final concentration Quantity or Volume
10× PBS 1× 100 mL
Water n/a 900 mL
Total n/a 1,000 mL
14. 1× PBST
Reagent Final concentration Quantity or Volume
10× PBS 1× 100 mL
Tween-20 0.1% 1 mL
Water n/a 900 mL
Total n/a 1,000 mL
15. Blocking solution
Reagent Final concentration Quantity or Volume
LiCOR blocking solution 50% 50 mL
PBST 50% 50 mL
Total n/a 100 mL
16. 10% APS solution
Reagent Final concentration Quantity or Volume
APS 10% 1 g
Water n/a 10 mL
Total n/a 10 mL
Laboratory supplies
1. Falcon 15 mL centrifuge tube (CORNING, catalog number: 352096)
2. Falcon 50 mL centrifuge tube (CORNING, catalog number: 352070)
3. Ultra-high performance centrifuge tubes (VWR, 525-1085)
4. SurPhob tips EcoReload, 1,250 μL (Biozym, catalog number: VT0174)
5. SurPhob tips EcoReload, 200 μL (Biozym, catalog number: VT0144)
6. SurPhob tips EcoReload, 10 μL (Biozym, catalog number: VT0104)
7. Transferpette S pipette 100–1,000 μL (BRAND, catalog number: BR705880)
8. Transferpette S pipette 20–200 μL (BRAND, catalog number: BR705878)
9. Transferpette S pipette 0.1–2.5 μL (BRAND, catalog number: BR705869)
10. Reaction tubes 1.5 mL (Biozym, catalog number: 710310)
11. 96-well flat bottom plate (Anicrin, catalog number: M09600P0)
12. Ice buckets for cooling steps
Equipment
1. Water bath (Lauda, model: Hydro H 20 S)
2. Centrifuge (Eppendorf, model: 5910 Ri, rotor: FA-6x50)
3. Odyssey Fc imaging system (Li-COR Odyssey Fc, model: 2800)
4. Homogenizer (Heidolf, model: Hei-TORQUE core)
5. Tissue grind pestle (Kimble Chase, catalog number: 885481-0023)
6. Mini-Protean Tetra cell casting modules (Bio-Rad, catalog number: 1658050)
7. Mini-Protean Tetra cell 4-gel system (Bio-Rad, catalog number: 1658004)
8. Criterion Blotter with wire electrodes (Bio-Rad, catalog number: 1704071)
9. Thermal shaker (VWR, model: Thermal shake lite, catalog number: 460-0249P)
10. SpectroStar Nano V5.70 (BMG Labtech)
Software and datasets
Image studio (Li-COR, Version 5.2)
MARS data analysis software (BMG Labtech, Version 3.42)
Procedure
A. Crude synaptosome preparation
Note: Ensure to maintain ice-cold conditions throughout all steps.
1. Sacrifice two mice by cervical dislocation and immediately dissect the brains.
Note: Treat each brain separately in different tubes for the following steps.
2. Place the brain in a pre-chilled tissue grinding pestle and add 8 mL of ice-cold sucrose buffer.
3. Homogenize at 900 rpm using 12 up-and-down strokes with the homogenizer.
4. Transfer the homogenate into a 15 mL centrifuge tube.
5. Spin the homogenate at 900× g for 10 min at 4 °C to remove large debris.
6. Transfer the supernatant (S1) into a 15 mL centrifuge tube and discard the pellet (P1).
7. Spin the collected supernatant (S1) in a centrifuge at 10,000× g for 15 min at 4 °C.
8. Discard the supernatant (S2) and resuspend the pellet (P2) in 2 mL of ice-cold sucrose buffer by gently pipetting up and down 3–4 times.
9. Add 6 mL of ice-cold sucrose buffer and gently invert to mix.
10. Spin the sample at 15,000× g for 15 min at 4 °C.
11. Discard the supernatant (S2’) and resuspend the pellet (P2’: crude synaptosomes) in 2 mL of sucrose buffer (see graphical overview for a schematic representation of synaptosomes preparation).
12. Combine the crude synaptosomes obtained from two brains, 4 mL in total, in a single 15 mL tube.
B. Protein estimation (Bradford assay)
1. Mix the Bradford reagent well before use and allow it to come to room temperature.
2. Prepare protein standards using a stock solution of 2 mg/mL BSA in sucrose buffer as mentioned in Table 1.
Table 1. Dilution scheme for protein standards using 2 mg/mL of BSA in sucrose buffer.
Vial Sucrose buffer Volume and source of BSA Final BSA concentration
A 0 300 μL of stock 2 mg/mL
B 125 μL 370 μL of stock 1.5 mg/mL
C 325 μL 325 μL of stock 1 mg/mL
D 175 μL 175 μL of vial B dilution 0.75 mg/mL
E 325 μL 325 μL of vial C dilution 0.5 mg/mL
F 325 μL 325 μL of vial E dilution 0.25 mg/mL
G 400 μL 0 0 mg/mL (blank)
3. Add 200 μL of Bradford reagent to a 96-well plate, add 2 μL of your protein standards or your sample (crude synaptosomes) to the reagent, and mix.
Note: Perform each measurement in triplicate and ensure that your actual sample concentration falls within the linear range of your standard curve, as other concentrations may lead to a nonlinear response.
4. Incubate for 5 min at room temperature in the dark.
5. Measure absorbance at 595 nm using the SpectroStar Nano V5.70.
6. Determine protein concentration using the MARS data analysis software.
Note: The usual protein concentration obtained from a single P60 mouse brain is 2–3 mg/mL (if the synaptosome yield is lower than expected, please refer to the troubleshooting section).
C. Proteolytic digestion procedure
1. For the untreated sample, add 5 mg of crude synaptosomes in a final volume of 10 mL of sucrose buffer without adding trypsin.
2. To initiate proteolytic digestion and to reach a final protein–protease ratio of 100:1 (see troubleshooting section), add 500 µL of trypsin solution (0.1 mg/mL) to 5 mg of crude synaptosomes in a final volume of 10 mL of sucrose buffer (see troubleshooting section).
3. Incubate the untreated sample and the protein–protease mixture for 10 min at 30 °C in a water bath, with occasional gentle inversion (no vortexing).
4. Centrifuge the samples at 8,700× g for 3 min at 4 °C.
5. Quickly resuspend the pellets in 1 mL of sucrose buffer containing 200 μL of 6× loading sample buffer to stop protease activity and in order to reach a final concentration of 5 µg/µL.
6. Boil the samples at 95 °C for 10 min.
7. The samples are now ready for SDS-PAGE to determine the subcellular localization of your protein of interest.
Pause point: Samples can be stored at -20 °C, and the following steps can be performed the day after. In our hands, samples can be stored at -20 °C for three months without degradation.
D. SDS gel casting and running
1. Clean the glass plates thoroughly and insert them into the casting frame.
2. Prepare the 10% separating gel solution (Table 2) in a 15 mL tube and swirl the solution gently.
Table 2. Recipe for 10% separating gel
Reagent Quantity or Volume
Water 3 mL
4× separating buffer 1.875 mL
Acrylamide/bis-acrylamide 2.5 mL
10% APS solution 150 μL
TEMED 15 μL
3. Pipette the separating gel solution between the glass plates, leaving 2 cm at the top.
4. Overlay the separating gel with isopropanol, making the separating gel regular and linear.
5. Once set, pour off completely the isopropanol and let it dry for 5 min.
6. Prepare the stacking gel solution (Table 3) in a 15 mL tube and swirl the solution gently.
Table 3. Recipe for 3.8% stacking gel
Reagent Quantity or Volume
Water 1.625 mL
4× stacking buffer 0.625 mL
Acrylamide/bis-acrylamide 0.333 mL
10% APS solution 75 μL
TEMED 7.5 μL
7. Pipette the stacking gel solution into the gap between the glass plates, layering it on top of the set separating gel.
8. Quickly insert the comb, ensuring to avoid any air bubbles, and allow it to sit for 10 min to set.
9. Carefully remove the comb once the stacking gel is fully set.
10. Take the gel out of the casting stand and insert it into the running cell buffer dam.
11. Pour 1× running buffer into the inner and outer chambers of the gel running apparatus and remove the comb.
12. Load the untreated and trypsin-treated samples (20–30 μg) into the wells along with a protein ladder/marker for size reference.
13. Run the gel at 80 V for the staking phase (approximately 40 min) and then increase to 130 V for the resolving phase, running until the dye front reaches the bottom of the gel.
E. Western blotting
1. Prepare the transfer materials by soaking the sponge, blotting paper, and nitrocellulose (NC) membrane in cold 1× transfer buffer.
2. Assemble the sandwich for the transfer in the following order: sponge—blotting paper—NC membrane—blotting paper—sponge.
3. Use a roller to gently remove any air bubbles between the layers for even transfer.
4. Fill the transfer chamber with cold 1× transfer buffer and insert the cool pack to maintain the temperature.
5. Insert the sandwich into the transfer chamber and run the transfer at 110 V for 100 min in a cold room or with the chamber on ice to prevent overheating.
F. Membrane staining
1. Carefully remove the NC membrane after the transfer and immediately stain it with Ponceau S staining solution for a few minutes to visualize the protein bands.
2. Rinse the membrane with water and take an image to record the transferred protein bands (see Figure 1).
3. Wash the membrane several times with 1× PBST until all traces of Ponceau S stain are gone.
4. Block the membrane by incubating it in blocking solution for 30–60 min to prevent nonspecific antibody binding.
5. While the membrane is blocking, prepare the primary antibody solution in a 15 mL tube using the blocking solution at the required dilution (see Table 4).
Table 4. Antibodies dilutions used in Figure 1
Antibody Dilution used Source
GluR1 1:1,000 Mouse
GluR2 1:1,000 Mouse
GluN2A 1:200 Rabbit
GluN2B 1:200 Rabbit
Synaptotagmin1 1:500 Rabbit
Synaptophysin1 1:500 Mouse
SNAP-25 1:500 Mouse
6. Transfer the membrane into the tube containing the primary antibody solution and incubate overnight at 4 °C on a rotator to ensure uniform binding.
7. The following day, wash the membrane with 1× PBST three times for 10 min each to remove any unbound primary antibodies.
8. Prepare the secondary antibody (mouse or rabbit, depending on the primary antibody used) solution at the dilution of 1:10,000 in 1× PBST.
9. Incubate the membrane with the secondary antibody for 45 min to 1 h at room temperature.
10. Wash the membrane with 1× PBST three times for 10 min each and then twice with 1× PBS to remove unbound secondary antibodies.
11. Visualize the membrane using the LiCOR imaging system to detect the protein of interest (see Figure 1) (refer to the troubleshooting section if variability in the proteolytic cleavage of postsynaptic proteins, unexpected degradation, or digestion of presynaptic proteins is observed).
Figure 1. Tryptic digest of synaptosomes reveals subcellular localization of synaptic proteins. A. Schematic representation of synaptosomes showing the presynaptic terminal resealed into an enclosed compartment, protecting presynaptic proteins from trypsin proteolysis, while postsynaptic proteins remain susceptible to tryptic digestion. B. Synaptosomes are either left untreated or incubated with trypsin, followed by immunoblot analysis. Presynaptic proteins, such as synaptotagmin1 (Syt1), SNAP-25, and synaptophysin1 (Syp), are protected from proteolysis. In contrast, postsynaptic proteins, including the AMPAR subunits GluA1 and GluA2, as well as the NMDAR subunits GluN2A and GluN2B, are sensitive to tryptic digestion, indicating their postsynaptic localization. C. Example of a Ponceau S staining of synaptosomes, either untreated or incubated with trypsin for 10 min at 30 °C. Please note that in the trypsin-treated sample, only postsynaptic proteins are cleaved. Therefore, there should not be a significant difference from the untreated sample. If many bands disappear, this may indicate over-digestion (see troubleshooting section).
Data analysis
The protocol essentially follows the workflow of the original synaptosome cleavage assay developed by Boyken et al. [1]. For data analysis in our publications [18–19], bands corresponding to synaptic proteins were quantified using the Empiria Studio Software package (LI-COR Biosciences). Band intensities of trypsin-treated samples were normalized to the untreated control, which is set to 100%, to calculate the relative abundance of each protein. A minimum of three biological replicates (n = 3) are recommended to ensure reproducibility, with data plotted as mean ± SEM. Statistical comparisons between groups (treated vs. untreated samples) can be performed using a one-sample t-test, comparing the relative abundance of trypsin-treated proteins to the hypothetical value of 100% from the untreated control. A p-value < 0.05 is considered significant. Examples of data analysis, quantification, and representation can be found in Figure 1B–C from [18] and Figure 4A from [19].
Validation of protocol
To our knowledge, this protocol or parts of it has been used and validated in the following research articles:
Boyken et al. [1]. Molecular Profiling of Synaptic Vesicle Docking Sites Reveals Novel Proteins but Few Differences between Glutamatergic and GABAergic Synapses. Neuron (Figure 1, panel B; Figure 2, panel A).
The protocol was originally developed for the isolation of a fraction highly enriched in synaptic vesicles docked to active zone. Mild proteolysis of synaptosomes was used to dissociate the presynaptic from the postsynaptic membrane.
Awasthi et al. [17]. Synaptotagmin-3 drives AMPA receptor endocytosis, depression of synapse strength, and forgetting. Science (Figure 1, panel E and F).
The protocol was used to verify the subcellular localization of synaptotagmin-3 (Syt-3). Awasthi et al. demonstrated that presynaptic proteins (such as synapsin, synaptobrevin-2, and Rab3a) were protected from trypsin cleavage, while postsynaptic proteins (such as Homer, GluA1, and PSD95), including Syt3, were cleaved.
Azarnia Tehran et al. [18]. Selective endocytosis of Ca2+-permeable AMPARs by the Alzheimer’s disease risk factor CALM bidirectionally controls synaptic plasticity. Science Advances (Figure 1, panel B and C).
The protocol was used to reveal the subcellular localization of the endocytic adaptor clathrin assembly lymphoid myeloid leukemia protein (CALM). We consistently found that while presynaptic proteins (such as SNAP-25, synaptophysin, and Rab3a) were protected from proteolysis, CALM was sensitive to trypsin, indicative of a large postsynaptic pool.
Li et al. [20]. P85S6K sustains synaptic GluA1 to ameliorate cognitive deficits in Alzheimer’s disease. Translational Neurodegeneration (Figure 1, panel C).
The protocol was used to verify the subcellular localization of p85S6K. As expected, presynaptic proteins (such as synaptophysin) were protected, while postsynaptic proteins (such as GluA1 and PSD95) were cleaved. Li et al. found that p85S6K, in both total and phosphorylated forms, was also sensitive to tryptic digestion, revealing its postsynaptic localization.
Vollweiter et al. [19]. Intersectin deficiency impairs cortico-striatal neurotransmission and causes obsessive–compulsive behaviors in mice. Proc Natl Acad Sci USA (Figure 4, panel A).
The protocol was used to verify the subcellular localization of intersectin1 (ITSN1). We found that ITSN1 is equally distributed between presynaptic and postsynaptic sites.
General notes and troubleshooting
General notes
1. This protocol requires basic knowledge of laboratory techniques and minimal animal surgical procedures.
2. To ensure optimal preservation of protein integrity during synaptosome isolation, it is critical that all procedures involving the preparation of synaptosomes are carried out on ice or at 4 °C. Always pre-chill your reagents and equipment (e.g., centrifuges) before starting the experiment.
3. In our publications, we used C57BL/6J mice, both male and female, above P60, at which point synaptogenesis and synaptic pruning are largely completed, and synaptic architecture is fully developed. The protocol should work for other strains and genetic backgrounds, with minimal adjustments. If the goal of the experiment is to assess developmental changes, using younger mice could provide valuable insights. However, some adjustments are recommended. For instance, approximately 3–4 times the number of brains may be needed to achieve similar protein yields. The same considerations apply to synaptosomes derived from specific mouse brain regions, which may be relevant in neurodegenerative diseases where only particular brain regions are affected, or from cultured neurons. Exact scaling should be determined experimentally when tissue availability is limited. For example, preliminary testing with different volumes is recommended to ensure effective synaptosome isolation.
4. The material obtained from two mouse brains is sufficient for both undigested and digested samples and can be used to check several known presynaptic and postsynaptic proteins as control, along with the protein(s) of interest via western blot. Since 5 mg of crude synaptosomes is enough for one experimental point, using two mouse brains also provides enough material to identify the optimal trypsin concentration (see troubleshooting section). However, if the optimal trypsin concentration is already known and fewer proteins need to be analyzed, the experiment can be scaled down to use a single mouse brain. In any case, we recommend verifying synaptosome integrity and confirming experimental success by checking at least a couple of known presynaptic and postsynaptic markers as control.
5. Due to trypsin digestion, we often observe bands at a lower molecular weight than the full-length postsynaptic protein of interest. This occurs because the antibody may also recognize peptide fragments generated by trypsin cleavage.
6. If proteins are partially cleaved (50%–70% remains), this may indicate that the protein is present at both presynaptic and postsynaptic ends, as seen with CALM and intersectin1 [18–19]. Nonetheless, we recommend combining results from the synaptosome trypsin cleavage assay with STED microscopy staining, using both presynaptic and postsynaptic markers. The combination of these methodologies will increase confidence in determining the sub-localization of your protein(s) of interest.
7. This protocol has been routinely used in our labs to assess the presynaptic and postsynaptic localization of various proteins by western blotting. However, it also has potential applications in combination with mass spectrometry. For example, the supernatant of trypsinized synaptosomes can be precipitated using chloroform–methanol precipitation, followed by further digestion with LysC and trypsin overnight at 37 °C, and then analyzed via mass spectrometry to identify postsynaptic proteins in high-throughput manner. Similarly, the pellet from trypsinized synaptosomes can be quickly lysed in lysis buffer (100mM Tris, 1% sodium deoxycholate, 10mM TCEP, and 12mM 2-chloroacetamide) by repeated sonication, followed by digestion with LysC and trypsin overnight at 37 °C to identify presynaptic proteins. In both cases, the resulting dry peptides can be stored at -80 °C.
8. Based on our experience, postsynaptic proteins (e.g., GluA1) show an optimal cleavage range of 90%, with an acceptable range of 80%–90%, meaning that the remaining protein after trypsin treatment should be 0%–20%. In contrast, presynaptic proteins (e.g., Syt1) have an optimal cleavage range of 0%–10%, with an acceptable extended range of 10%–20%, so that the remaining protein amount should be 80%–100%. When proteins show intermediate cleavage levels (40%–70%), these proteins are inferred to localize to both presynaptic and postsynaptic sites.
Troubleshooting
Problem 1: Low yield of synaptosomes.
Possible cause: Low yield of synaptosomes can result from insufficient tissue homogenization, improper centrifugation, or poor buffer quality.
Solution: We recommend using a mechanical homogenizer, as it provides the consistent shear force necessary for effective brain disruption while preserving synaptic structures. However, we noticed that other published protocols have successfully used a Dounce homogenizer, with 15–20 strokes using a loose pestle to achieve comparable disruption. We also recommend checking centrifugation speed and time, ensuring that tubes are properly balanced. Finally, fresh buffers should be prepared before starting synaptosome preparation.
Problem 2: Variability in proteolytic digestion of postsynaptic proteins.
Possible cause: Partial resistance of postsynaptic proteins to proteolytic degradation due to the densely packed postsynaptic network.
Solution: In the original publication, Boyken et al. [1] observed that certain postsynaptic proteins, such as PSD95 and Homer1, were not degraded, suggesting partial resistance of the postsynaptic density network to proteolytic digestion. In our experience, Homer1 is resistant to trypsin cleavage, while PSD95 is cleaved. However, in the study by Awasthi et al. [17], both PSD95 and Homer1 were cleaved after 10 min of trypsin digestion. This variability may result from differences in synaptosome preparation and/or the antibodies used for western blot detection. We would like to emphasize to readers that although this assay can be used in a high-throughput manner to screen various presynaptic and postsynaptic proteins, their localization should be confirmed with additional methods, such as super-resolution microscopy (e.g., STED), to unequivocally verify proper localization.
Problem 3: Unexpected degradation of proteins during synaptosome preparation.
Possible cause: If protein degradation is observed even in the untreated sample, where trypsin was not added, this suggests degradation due to endogenous cellular proteases released during preparation. Since synaptosome fractionation is performed without protease inhibitors to avoid interference with trypsin activity, we highly recommend conducting all steps at 4 °C and pre-chilling all equipment.
Problem 4: Digestion of presynaptic proteins.
Possible cause: Prolonged digestion time or excessive trypsin digestion.
Solution 1: This protocol relies entirely on the integrity of synaptosomes. If the purified synaptosomes are damaged or broken, trypsin will also access presynaptic proteins. To address this issue, we recommend reducing variability in synaptosome preparations across experiments by monitoring synaptosome yield and purity at intermediate steps (e.g., after each centrifugation). It is especially important to ensure that all materials (e.g., homogenizer) used during synaptosome preparation are detergent-free, as any traces of detergent can compromise synaptosome integrity and thereby affect the final results.
Solution 2: Extended incubation can lead to over-digestion, resulting in degradation of presynaptic proteins. Ensure that trypsin digestion is carefully monitored, carried out at 30 °C for 10 min, followed by immediate centrifugation and the addition of 1 mL sucrose buffer containing 200 μL of 6× loading sample buffer to stop the reaction.
Solution 3: Trypsin potency can vary between different batches. Therefore, it is essential to test each new batch of trypsin before use. Perform a trypsin cleavage assay using various protein-to-protease ratios, such as 50:1, 100:1, or 250:1, to determine the optimal digestion conditions. Select the ratio that provides effective postsynaptic cleavage without leading to over-digestion of presynaptic proteins For example, the old batch of trypsin used in Azarnia Tehran et al. [18] was used with a protein-to-protease ratio of 100:1. In Vollweiter et al. [19], the new batch of trypsin worked best with a protein-to-protease ratio of 50:1 (see Figure 2). With this new batch of trypsin, using a higher trypsin concentration led to cleavage of presynaptic proteins as well (e.g., synaptotagmin1, Syt1).
Figure 2. Determination of optimal protein-to-protease ratio for the trypsin cleavage assay. Representative immunoblot showing the effects of varying protein-to-protease ratios (50:1, 100:1, and 250:1) on the cleavage of the postsynaptic protein GluA1 and the presynaptic protein synaptotagmin1 (Syt1). Synaptosomes were left untreated or were treated with increasing trypsin concentrations. GluA1 is sensitive to trypsin digestion across all conditions. In contrast, Syt1 is resistant at the 50:1 ratio but becomes partially cleaved at higher trypsin concentrations, demonstrating the importance of optimizing the trypsin concentration to preserve presynaptic proteins.
Problem 5: Weak or absent protein bands in western blot.
Possible cause: Insufficient protein loading.
Solution: We typically load 20–30 μg of protein per well. However, depending on the antibody used, this amount might be insufficient for detection. To address this, optimize the protein loading concentration and adjust the primary and secondary antibody dilutions. Experimenting with different concentrations and dilutions can help enhance the signal and achieve better detection.
Acknowledgments
This protocol resulted from the adaptation of the original research article from Boyken et al. (DOI: 10.1016/j.neuron.2013.02.027). This protocol was used by the authors in Azarnia Tehran et al. [18] and Vollweiter et al. [19]. This work was supported by a Klaus Tschira Boost Fund from the Klaus Tschira Stiftung to D.A.T.
BioRender.com was used for the graphical overview and Figure 1.
Competing interests
D.A.T. is an Assistant Editor for Bio-protocol but did not participate in the editorial and peer review process of this article, except as an author. The authors declare no other conflict of interest.
Ethical considerations
All animal experiments performed in Azarnia Tehran et al. [18] and Vollweiter et al. [19] were reviewed and approved by the ethics committee of the “Landesamt für Gesundheit und Soziales” (LAGeSo) Berlin and were conducted according to the committee’s guidelines.
References
Boyken, J., Grønborg, M., Riedel, D., Urlaub, H., Jahn, R. and Chua, J. J. E. (2013). Molecular Profiling of Synaptic Vesicle Docking Sites Reveals Novel Proteins but Few Differences between Glutamatergic and GABAergic Synapses. Neuron. 78(2): 285–297. https://doi.org/10.1016/j.neuron.2013.02.027
Südhof, T. C. (2013). Neurotransmitter Release: The Last Millisecond in the Life of a Synaptic Vesicle. Neuron. 80(3): 675–690. https://doi.org/10.1016/j.neuron.2013.10.022
Haucke, V., Neher, E. and Sigrist, S. J. (2011). Protein scaffolds in the coupling of synaptic exocytosis and endocytosis. Nat Rev Neurosci. 12(3): 127–138. https://doi.org/10.1038/nrn2948
Rizzoli, S. O. (2014). Synaptic vesicle recycling: steps and principles. EMBO J. 33(8): 788–822. https://doi.org/10.1002/embj.201386357
Kononenko, N. L. and Haucke, V. (2015). Molecular Mechanisms of Presynaptic Membrane Retrieval and Synaptic Vesicle Reformation. Neuron. 85(3): 484–496. https://doi.org/10.1016/j.neuron.2014.12.016
Azarnia Tehran, D., López-Hernández, T. and Maritzen, T. (2019). Endocytic Adaptor Proteins in Health and Disease: Lessons from Model Organisms and Human Mutations. Cells. 8(11): 1345. https://doi.org/10.3390/cells8111345
Azarnia Tehran, D. and Maritzen, T. (2022). Endocytic proteins: An expanding repertoire of presynaptic functions. Curr Opin Neurobiol. 73: 102519. https://doi.org/10.1016/j.conb.2022.01.004
Li, Y. C. and Kavalali, E. T. (2017). Synaptic Vesicle-Recycling Machinery Components as Potential Therapeutic Targets. Pharmacol Res. 69(2): 141–160. https://doi.org/10.1124/pr.116.013342
Michetti, C., Falace, A., Benfenati, F. and Fassio, A. (2022). Synaptic genes and neurodevelopmental disorders: From molecular mechanisms to developmental strategies of behavioral testing. Neurobiol Dis. 173: 105856. https://doi.org/10.1016/j.nbd.2022.105856
Timalsina, B., Lee, S. and Kaang, B. K. (2024). Advances in the labelling and selective manipulation of synapses. Nat Rev Neurosci. 25(10): 668–687. https://doi.org/10.1038/s41583-024-00851-9
Xu, Y., Song, X., Wang, D., Wang, Y., Li, P. and Li, J. (2021). Proteomic insights into synaptic signaling in the brain: the past, present and future. Mol Brain. 14(1): 37. https://doi.org/10.1186/s13041-021-00750-5
Hindley, N., Sanchez Avila, A. and Henstridge, C. (2023). Bringing synapses into focus: Recent advances in synaptic imaging and mass-spectrometry for studying synaptopathy. Front Synaptic Neurosci. 15: e1130198. https://doi.org/10.3389/fnsyn.2023.1130198
Marcassa, G., Dascenco, D. and de Wit, J. (2023). Proteomics-based synapse characterization: From proteins to circuits. Curr Opin Neurobiol. 79: 102690. https://doi.org/10.1016/j.conb.2023.102690
van Oostrum, M., Blok, T. M., Giandomenico, S. L., tom Dieck, S., Tushev, G., Fürst, N., Langer, J. D. and Schuman, E. M. (2023). The proteomic landscape of synaptic diversity across brain regions and cell types. Cell. 186(24): 5411–5427.e23. https://doi.org/10.1016/j.cell.2023.09.028
Arizono, M. and Nägerl, U. V. (2021). Deciphering the functional nano‐anatomy of the tripartite synapse using stimulated emission depletion microscopy. Glia. 70(4): 607–618. https://doi.org/10.1002/glia.24103
Yang, X. and Annaert, W. (2021). The Nanoscopic Organization of Synapse Structures: A Common Basis for Cell Communication. Membranes (Basel). 11(4): 248. https://doi.org/10.3390/membranes11040248
Awasthi, A., Ramachandran, B., Ahmed, S., Benito, E., Shinoda, Y., Nitzan, N., Heukamp, A., Rannio, S., Martens, H., Barth, J., et al. (2019). Synaptotagmin-3 drives AMPA receptor endocytosis, depression of synapse strength, and forgetting. Science (1979). 363(6422): eaav1483. https://doi.org/10.1126/science.aav1483
Azarnia Tehran, D., Kochlamazashvili, G., Pampaloni, N. P., Sposini, S., Shergill, J. K., Lehmann, M., Pashkova, N., Schmidt, C., Löwe, D., Napieczynska, H., et al. (2022). Selective endocytosis of Ca 2+ -permeable AMPARs by the Alzheimer’s disease risk factor CALM bidirectionally controls synaptic plasticity. Sci Adv. 8(21): eabl5032. https://doi.org/10.1126/sciadv.abl5032
Vollweiter, D., Shergill, J. K., Hilse, A., Kochlamazashvili, G., Koch, S. P., Mueller, S., Boehm-Sturm, P., Haucke, V. and Maritzen, T. (2023). Intersectin deficiency impairs cortico-striatal neurotransmission and causes obsessive–compulsive behaviors in mice. Proc Natl Acad Sci USA. 120(35): e2304323120. https://doi.org/10.1073/pnas.2304323120
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Article Information
Publication history
Received: Oct 8, 2024
Accepted: Nov 21, 2024
Available online: Dec 10, 2024
Published: Jan 20, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Neuroscience > Cellular mechanisms > Synaptic physiology
Biochemistry > Protein > Isolation and purification
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Flow-based In Vivo Method to Enumerate Translating Ribosomes and Translation Elongation Rate
MS Mina O. Seedhom
DD Devin Dersh
JY Jonathan W. Yewdell
Published: Vol 15, Iss 2, Jan 20, 2025
DOI: 10.21769/BioProtoc.5165 Views: 1025
Reviewed by: Chiara AmbrogioThirupugal GovindarajanIstvan Stadler
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Original Research Article:
The authors used this protocol in eLIFE Mar 2024
Abstract
Protein synthesis is by far the most energetically costly cellular process in rapidly dividing cells. Quantifying translating ribosomes in individual cells and their average mRNA transit rate is arduous. Quantitating assembled ribosomes in individual cells requires electron microscopy and does not indicate ribosome translation status. Measurement of average transit rates entails in vitro pulse-chase radiolabeling of isolated cells or ribosome profiling after ribosome runoff, which is expensive and extremely demanding technically. Here, we detail protocols based on ribosome-mediated nascent chain puromycylation, harringtonine to stall initiating ribosomes while allowing ribosome elongation to continue normally, and cycloheximide to freeze translating ribosomes in place. Each compound is delivered intravenously to mice in the appropriate order, and after ex vivo cell fixation and permeabilization, translating ribosome numbers and transit rates are measured by flow cytometry using a directly conjugated puromycin-specific antibody.
Key features
• Measure relative numbers of translating ribosomes in mixed single-cell preparations.
• Quantitate relative in vivo ribosome transit rates in mixed single-cell preparations.
• Detect ribosome stalling in vivo.
Keywords: Protein synthesis Puromycin Ribosome Nascent chain RiboPuroMycylation (RPM) Ribosome transit analysis (RTA)
Graphical overview
Background
Proteins are the most abundant macromolecule by mass and copy number in nearly all cell types in varied proliferation states [1,2]. Understanding proteostasis, the net outcome of ribosomal synthesis, folding, secretion/release, and degradation, requires an accurate accounting of ribosome numbers and translation rates.
The protocol we describe here is based on puromycin (PMY), a remarkable aminonucleoside antibiotic that interferes with protein synthesis in all forms of life. By mimicking an aminoacylated tyrosyl tRNA, PMY is incorporated by ribosome catalysis into nascent chains, where it terminates elongation as it lacks a COOH terminus that can form a peptide bond [3,4]. This reaction, termed puromycylation, occurs spontaneously, even at 0 °C. It has been utilized (in conjunction with various protein translation modulators) by our laboratory to uncover evidence of nuclear translation [5], antigen-independent cytokine-induced division of memory-like T cells in the bone marrow of mice after virus infection [6], and stalled ribosomes in neurons and immune cells [7,8], along with many discoveries by other labs [9].
We recently reported protocols that measure the numbers of translating ribosomes and ribosome transit rates in mice based on intravenous (IV) delivery of protein synthesis modulators and flow cytometry–based detection of puromycylated nascent chains using a fluorescently labeled anti-puromycin antibody (Ab) [8]. Herein, we describe these protocols in detail.
Materials and reagents
Biological materials
1. C57BL/6J mice, male or female (strain number: 00664)
2. BD PharmingenTM purified rat anti-mouse CD16/CD32 (mouse BD Fc Block™) (BD Bioscience, catalog number: 553141)
Reagents
1. Puromycin dihydrochloride from Streptomyces alboniger (PMY) (Sigma-Aldrich, catalog number: P8833-100MG)
2. Cycloheximide (CHX) (EMD Millipore, catalog number: 239764, 100 mg)
3. Methanol (certified ACS) (Fisher Chemical, catalog number: A412-4)
4. Harringtonine (HAR) (Santa Cruz Biotech, catalog number: sc-204771A)
5. RPMI 1640 medium, GlutaMAXTM supplement (Thermo Fisher Scientific, catalog number: 61870036)
6. DPBS, no calcium, no magnesium (Thermo Fisher Scientific, catalog number: 14190144)
7. Anti-puromycin Ab, produced from in-house-generated hybridoma (Developmental Studies Hybridoma Bank, catalog number: PMY-2A4)
8. ACK lysing buffer (Lonza, catalog number: A1049201)
9. Trypan Blue solution, 0.4% (Thermo Fisher Scientific, catalog number: 15250061)
10. Fujifilm Wako chemicals USA digitonin 1 g (Fisher Scientific, catalog number: NC0141730)
11. Paraformaldehyde, 16% w/v aq. soln., methanol free (Thermo Fisher Scientific, catalog number: 043368.9M)
12. Fluorescent Protein Labeling kits (Thermo Fisher Scientific, catalog number: A10235)
13. Ethidium monoazide bromide (EMA) (Thermo Fisher Scientific, catalog number: E1374)
14. Bovine serum albumin (BSA) (Millipore Sigma, catalog number: A7888-10G)
15. Sodium azide (Millipore Sigma, catalog number: 71290)
16. Ethyl alcohol, pure (Millipore Sigma, catalog number: 459836-100ML)
17. Fetal bovine serum (FBS) (Atlanta Biologicals, catalog number: S10350)
Solutions
1. RPMI with 7.5% fetal bovine serum (see Recipes)
2. Puromycin (PMY), cycloheximide (CHX), and harringtonine (HAR) solution for IV injections (1 mL) (see Recipes)
3. Stock CHX solution (see Recipes)
4. Stock HAR solution (see Recipes)
5. Stock PMY solution (see Recipes)
6. HAR/PMY/CHX solution for intravenous injection (see Recipes)
7. HAR solution for intravenous injection (see Recipes)
8. Fluorescence-activated cell sorting (FACS) buffer (see Recipes)
9. Stock EMA solution (see Recipes)
10. Stock digitonin solution (see Recipes)
11. Fixation and permeabilization buffer (see Recipes)
Recipes
1. RPMI 1640 with 7.5% FBS
500 mL of RPMI 1640 with GlutaMAX
37.5 mL of FBS
Store at 4 °C.
2. Puromycin (PMY), cycloheximide (CHX), and harringtonine (HAR) solution for IV injections (1 mL)
1 mg of PMY, 100 μg of HAR, and 0.34 mg of CHX per mouse; 100 μL per mouse; make fresh from stocks.
20 μL of HAR stock solution
200 μL of PMY stock solution
170 μL of CHX stock solution
610 μL of PBS
Warm to 37 °C before use to allow PMY to fully solubilize, then keep at room temperature for up to 5 h.
3. Harringtonine (HAR) solution for IV infections (1 mL)
100 μg of HAR per mouse, 100 μL per mouse, make fresh from stocks.
20 μL of HAR stock solution
980 μL of PBS
Once made, use immediately. Can remain at room temperature for ~5 h.
4. CHX stock solution in 50% ethanol (20 mg/mL)
100 mg of CHX
5 mL of 50% ethanol, 50% water
Aliquot and store at -20 °C.
5. PMY stock solution in PBS (50 mg/mL)
100 mg of puromycin (PMY)
2 mL of PBS
Warm to 37 °C to solubilize, aliquot (we usually use 1 mL), and store at -20 °C.
6. Harringtonine (HAR) stock solution in methanol (50 mg/mL)
10 mg of HAR
200 μL of methanol
Aliquot and store at -20 °C.
7. 1% digitonin stock solution in DMSO
1 g of digitonin
100 mL of DMSO
Aliquot and store at -20 °C.
8. Fixation/permeabilization buffer (1% paraformaldehyde, 0.0075% digitonin)
9.3 mL of PBS
625 μL of 16% paraformaldehyde
75 μL of 1% digitonin
Make fresh and keep on ice.
9. FACS buffer (0.2% sodium azide, 1 mg/mL bovine serum albumin)
1 g of sodium azide
500 mg of BSA
500 mL of PBS
Store at 4 °C.
10. EMA stock solution (5 mg/mL)
5 mg of EMA
1 mL of DMSO
Aliquot and store at -20 °C.
11.Ethidium monoazide solution for staining (10 μg/mL)
20 μL of EMA stock solution
10 mL of PBS
Make fresh and keep on ice.
Laboratory supplies
1. FalconTM 15 mL conical polypropylene centrifuge tubes (Fisher Scientific, catalog number: 14-959-49B)
2. Corning® Primaria TM 60 mm × 15 mm standard cell culture dish (Corning, catalog number: 353802)
3. Microscope slides, glass, 25 × 75 mm, 90° ground edges, frosted, 1 end, both sides (Globe Scientific, catalog number: 1308)
4. Falcon® 70 μm cell strainer, white, sterile, individually packaged, 50 per case (Fisher Scientific, catalog number: 22-363-548)
5. Cellometer SD025 slides, box of 75 slides (Revvity, catalog number: CHT4-SD025-002)
6. Corning® 96-well clear round bottom TC-treated microplate (Corning, product number: 3799)
7. BD EclipseTM needle 27 G × 1/2 in. with detachable 1 mL BD Luer-LokTM syringe (BD Biosciences, GTIN number: 00382903057894)
Equipment
1. Cellometer X2 fluorescent viability counter (Nexcelom, catalog number: CMT-X2-S150)
2. Tailveiner restrainer for mice (Braintree Scientific, Inc., SKU: TV-150)
3. Infrared heating and drying lamp, table model (Walter Stern, VWR, Avantor, catalog number: 36547-009)
4. FisherbrandTM accuSpin Max 1.6 L benchtop centrifuge (Fisher Scientific, catalog number: 75-883-78)
5. LSR Fortessa X-20 (BD Bioscience, catalog number: 50165)
Software and datasets
1. Microsoft Excel for Mac, version 16.85
2. GraphPad Prism 10 for MacOS
3. FlowJo 10.10.0 for Mac OS X
Procedure
A. Intravenous injections of reagents necessary for in vivo RiboPuroMycylation (RPM)
Note 1: All in vivo procedures require approval by your institutional Animal Care and Use Committee (ACUC).
Note 2: The in vivo RPM procedure has been successfully performed on lymph nodes, spleens, and thymi from mice, although a similar procedure has been performed in vitro on other organs such as fetal liver [10].
Note 3: For in vivo RPM and RTA procedures, we have used female and male mice aged 1.5–18 months. Mice were housed on a 12 h light/dark cycle with ad libitum access to normal mouse chow and water. The procedure was developed using C57BL/6J (B6) mice, and we have performed it using other mice on the B6 and FVB/N background. We expect this protocol will work similarly in other mouse strains and animal models, with adjustments to drug dosage.
1. For the in vivo RPM assay, prepare a solution of 10 mg/mL PMY, 1 mg/mL HAR, and 3.4 mg/mL CHX in PBS and warm in a water bath to 37 °C to allow PMY to completely solubilize (see Recipes). Prepare approximately 20% extra of this solution, as fluid loss during injections will occur.
2. Prepare a solution of 1 mg/mL of HAR in PBS (see Recipes). Prepare approximately 20% extra of this solution, as fluid loss during injections will occur.
3. Warm mice for 5–10 min using an ACUC-approved heat lamp to dilate tail veins. Mice are considered sufficiently warm for injections when their activity within the cage is increased.
4. Restrain warmed mice in a mouse holder and intravenously inject the tail vein, using a 27G × ½ needle attached to a 1 mL syringe, with 100 μL of the HAR/CHX/PMY solution, waiting 5 min for the maximum signal. To determine the background RPM signal not related to active protein synthesis, inject mice with 100 μL of the HAR solution, wait 15 min for ribosomes to finish translating mRNAs, and then inject 100 μL of the HAR/CHX/PMY solution and wait 5 min. A third control is necessary, injecting vehicle without PMY to control for non-specific staining of the anti-PMY mAb.
5. After the indicated times, sacrifice mice by cervical dislocation and harvest organs of interest into 15 mL conical centrifuge tubes containing 3 mL of RPMI supplemented with 7.5% FBS on ice.
Figure 1. Depiction of the in vivo RiboPuroMycylation (RPM)-ribosome transit assay (RTA). In this example, CFSE-labeled OT-1 T cells (CFSE tracks cell division) are adoptively transferred into congenic mice (to allow tracking of donor cells in recipient mice), followed by infection with vaccinia virus expressing the SIINFEKL peptide (Kb-SIINFEKL activates OT-1 T cells). RPM-RTA is performed by intravenous injection of harringtonine (HAR) for different amounts of time followed by HAR, cycloheximide (to prevent leakiness from HAR inhibition alone), and puromycin. Spleens are harvested for RPM analysis on both endogenous and transferred T cells. Schematic designed with BioRender.
B. Intravenous injections of reagents necessary for in vivo RPM ribosome transit analysis (RTA)
Note: The timing of intravenous injections is critical. It is strongly suggested that this procedure be done by a two-person team: one person using a timer to record the time of injections and the other to perform the injections.
1. For in vivo RPM RTA (depiction of an experimental setup in Figure 1), prepare a solution of 10 mg/mL PMY, 1 mg/mL HAR, and 3.4 mg/mL CHX and warm in a water bath to 37 °C to allow PMY to completely solubilize (see Recipes). Prepare approximately 20% extra of this solution, as fluid loss during injections will occur.
2. Prepare a solution of 1 mg/mL HAR (see Recipes). Prepare approximately 20% extra of this solution, as fluid loss during injections will occur.
3. Warm mice for 5–10 min using an ACUC-approved heat lamp to dilate tail veins. Mice are considered sufficiently warm for injections when their activity within the cage is increased.
4. Restrain warmed mice in a mouse holder, intravenously inject the tail vein using a 27G × ½ needle attached to a 1 mL syringe with 100 μL of the HAR/CHX/PMY solution, and wait 5 min for maximum signal. To determine ribosome transit times, inject 100 μL of HAR and wait for 30 s, 1 min, 2 min, 4 min, or 10 min before injecting with 100 μL of the HAR/CHX/PMY solution. As above, a no-PMY control is needed.
5. After the indicated times, sacrifice mice by cervical dislocation and harvest organs of interest into 15 mL conical centrifuge tubes containing 3 mL of RPMI supplemented with 7.5% FBS on ice.
C. Preparation of single-cell samples for flow cytometry
Note 1: All steps below are performed at 4 °C with ice-cold solutions.
Note 2: We have stained anywhere from 1 to 6 million cells per well in a 96-well plate, with adjustments to antibody concentrations for surface and intracellular antigens.
1. Crush organs (we have used spleens, thymi, or lymph nodes) in 3 mL of RPMI between two frosted microscope slides in a 60 mm × 15 mm standard cell culture dish. Filter resultant single-cell suspensions through a 70 μm mesh screen back into a standard polyethylene 15 mL centrifuge tube.
2. Lyse red blood cells present in tissue samples by adding 6 mL of ACK lysing buffer directly to the single cell suspensions and mix by quickly rotating the tubes.
3. Centrifuge filtered single-cell suspensions at 350× g for 4 min at 4 °C and pour off supernatants. Disrupt cell pellets by tapping the bottom of the 15 mL conical centrifuge tubes quickly 3–4 times. Resuspend cells in 3 mL of cold RPMI supplemented with 7.5% FBS. Centrifuge at 350× g for 4 min at 4 °C, pour off supernatants, disrupt cell pellet, and add 3 mL of cold RPMI supplemented with 7.5% FBS. Refilter cell suspensions again through a 70 μm mesh screen.
4. Count resuspended cells with a Nexcelom Cellometer Vision using Trypan Blue (1:1 dilution) for live/dead cell discrimination as per the manufacturer’s instructions. After counting, centrifuge the cell suspensions in 15 mL conical centrifuge tubes at 350× g for 4 min at 4 °C, pour off supernatants, disrupt cell pellets, and resuspend cells in cold RPMI supplemented with 7.5% FBS at 10 million cells per milliliter. Add 200 μL (1–6 million cells per well can be plated; here, 2 million cells per well are plated) of the cell suspensions from each sample to a well of a 96-well polystyrene U-bottom plate.
D. Surface and intracellular stains for flow cytometry analysis
Note 1: Stains for flow cytometry are all performed on ice.
Note 2: Anti-puromycin Ab should be conjugated with the required fluorochrome ahead of time with a Life Technologies Protein Labeling kit as per the manufacturer’s instructions.
Note 3: The fluorescence signal from the fluorochrome-conjugated anti-puromycin Ab will dim over time. It is recommended that you only use the conjugated anti-PMY Ab for six months.
1. For single-color controls (SCC) and fluorescence-minus-one (FMO) controls, prepare a mixture of equal numbers of cells from each sample for each organ type isolated (all splenocytes in one mixture, all cells from thymi in a separate mixture). This mixture is made for appropriate flow cytometer setup and downstream analysis. New SCCs and FMOs should be prepared for each experiment. Settings on the flow cytometer may be reused, depending on the experience of the experimenter.
2. Mix Abs at proper dilutions in FACS buffer for SCCs, FMOs, and the full Ab mix to allow for 100 μL per sample in the 96-well plate. Centrifuge cells in the 96-well plate at 350× g for 4 min at 4 °C. Before dumping the supernatant [with one (!) quick smooth wrist flick into a biohazard bucket], check that solid cell pellets have formed at the bottom of the 96-well plate.
3. Resuspend cells in 200 μL of PBS using a 12-well multichannel pipette. Centrifuge the plate as above. Dump the supernatant and resuspend the appropriate FMOs, SCCs, and samples in 100 μL of a 10 μg/mL solution of EMA in PBS for flow-based live/dead cell discrimination (see Recipes). Place the plate in the dark on ice for 10 min. Next, expose the uncovered plate close to a fluorescent light with the lid off for an additional 10 min on ice. While exposure to a commercial fluorescent light is suitable for crosslinking EMA to DNA, a wavelength of 465–475 nm has been described to work in the PMA-LiteTM 2.0 LED Photolysis Device (Biotium, catalog number: E90006). Add 100 μL of PBS to each well, pipette up and down to mix, and centrifuge as above. After washing with 200 μL of PBS twice, perform surface staining.
4. Resuspend cells in 100 μL of FACS buffer with the 2.42G monoclonal antibody (Ab) to block Fc receptors (dilution as per manufacturer’s instructions) on ice for 10 min. Add 100 μL of the appropriate single Abs for SCCs, Ab mixes for FMOs, or full Ab mix for full stains, to cell surface antigens at appropriate dilutions for 30 min at 4 °C (a common Ab panel we use and the associated dilutions are listed in Table 1). Centrifuge as above and wash twice in 200 μL of PBS.
Table 1. Example flow cytometry panel
Antibody and fluorochrome Manufacturer Catalog number Dilution
BD Horizon BV786 hamster anti-mouse CD3e (clone 145-2-C11) BD Bioscience 564379 0.75 μL per test
BD Horizon BV510 rat anti-mouse CD4 (clone RM4-5) BD Bioscience 563106 0.75 μL per test
BD Horizon PE-CF594 rat anti-mouse CD8a (clone 53-6.7) BD Bioscience 562283 0.75 μL per test
BD Pharmingen PE mouse anti-mouse Vβ 5.1, 5.2 T-cell receptor (clone MR9-4) BD Bioscience 562086 0.75 μL per test
eBioscience PE-Cyanine7 rat anti-mouse CD19 (clone eBio1D3 (1D3) Invitrogen 25-0193-82 0.75 μL per test
eBioscience APC mouse anti-mouse CD45.1 (clone A20) Invitrogen 17-0453-82 0.75 μL per test
eBioscience mouse anti-mouse Super Bright 780 CD45.2 monoclonal Ab (clone 104) Invitrogen 78-0454-82 0.75 μL per test
5. Next, simultaneously fix and permeabilize cells in 100 μL of freshly made fixation/permeabilization buffer (fix/perm) (1% PFA, 0.0075% digitonin in PBS, see Recipes) for 20 min at 4 °C. Add 100 μL of PBS, centrifuge as above, and wash twice with PBS.
6. Stain for PMY after the fix/perm step using a 1:100 dilution of the conjugated anti-PMY Ab for 1 h (it could be overnight if more convenient). Add 100 μL of PBS, pipette up and down 6–8 times, and centrifuge the plate as above. Wash twice with 200 μL of PBS, resuspend cell pellets in FACS buffer, and analyze samples (samples should remain on ice during the run) with a flow cytometer. Samples may be stored in a dark refrigerator for up to a week prior to running on a flow cytometer. We use a BD LSRII or BD LSR Fortessa X-20 to run samples, but any flow cytometer with the appropriate lasers and filter sets may be used.
Data analysis
1. Analyze resulting data using FlowJo, Microsoft Excel, and GraphPad Prism software. For our analysis, we gated OT-1 CD8+ T cells by gating on singlets by FSCa and FSCw, lymphocytes by SSCa and FSCa, live cells by EMA-, T cells by CD3+CD19-, CD8+ T cells by CD8+CD4-, adoptively transferred cells by CD45.1+CD4-, and OT1 T cells by Vb5+CD45.2-, as shown in Figure 2a. For RPM values, we took the population of interest, in this case adoptively transferred OT-1 CD8+ T cells, and determined the mean fluorescence intensity (MFI) of PMY staining in mice treated with different inhibitors of either the CFSE+ population in uninfected mice or the CFSE- population in day 3 VACV-SIINFEKL-infected mice. We chose an OT1 T-cell experiment with CFSE labeling (CFSE tracks cell division, CFSE hi are undivided, CFSE low are divided) to distinguish the difference in protein synthesis in actively dividing vs. resting (non-dividing) T cells.
2. For the in vivo RPM assay, subtract the no-PMY MFI in your population of interest (OT1 T cells here) from the HAR/CHX/PMY-treated PMY MFI (maximum signal). Next, subtract the no-PMY PMY MFI signal from the 15-min HAR-treated and then the HAR/CHX/PMY PMY signal.
Note: Typically, only two no-PMY-treated mice are required for each manipulation, and the average of the RPM MFIs of the no-PMY-treated mice are taken, as the MFI for each no-PMY mouse should be very similar.
3. Next, subtract the no-PMY-subtracted 15-min HAR-treated and then HAR/CHX/PMY-treated PMY signal from the no-PMY-subtracted HAR/CHX/PMY PMY signal. This will be the relative number of ribosomes in the cell population analyzed (RPM value). If the 15-min HAR-treated and then HAR/CHX/PMY-treated runoff RPM signal is consistently higher than the no-PMY RPM signal, this could be an example of stalled translation, as we have seen in cultured un-activated lymphocytes in vitro [8] and has also been described in neurons [7].
4. For the in vivo RTA assays, the initial analysis is the same, but as the analysis is for different times after runoff, the resultant values are plotted in GraphPad Prism against the time of each individual HAR treatment, and a one-phase decay analysis is performed on the resultant graph. An example of graphed data normalized to the maximum signal from three different experiments is shown in Figure 2b. In this example, we examine translation elongation rates in both non-activated polyclonal T cells and activated OT1 transgenic T cells.
Figure 2. Example gating strategy and RiboPuroMycylation ribosome transit analysis data A. On the left: Gating on OT-1 T CD8+ T cells. Singlets by FSCa and FSCw, lymphocytes by SSCa and FSCa, live cells by EMA-, CD3+CD19-, CD8+ T cells by CD8+CD4-, adoptively transferred cells by CD45.1+CD4-, and OT1 T cells by Vb5+CD45.2-. On the right: RiboPuroMycylation (RPM) staining of divided (CFSE-, infected mice) or undivided (CFSE+, uninfected mice) OT-1 CD8+ T cells, with an example of harringtonine runoff after different harringtonine treatment times in CFSE- cells. B. RPM-Ribosome Transit Analysis (RTA) of adoptively transferred activated OT-1 T cells or un-activated (resting) host CD8+ T cells in mice infected for 2 or 3 days with VACV-SIINFEKL. Three to four independent experiments combined. Normalized by setting the corrected maximum RPM signal (no runoff) to 100.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article:
Seedhom et al. [8]. Paradoxical imbalance between activated lymphocyte protein synthesis capacity and rapid division rate. eLife.
General notes and troubleshooting
1. While extensively validated in lymphocytes and CFSE-labeled lymphocytes from the spleen, thymus, and lymph nodes in two different strains of mice (C57BL/6J and FVB/N) as well as RAG1ko OT1 TCR transgenic mice, this protocol relies on intravenous delivery of protein synthesis modifiers and detection reagents that work similarly in many mammals, so we expect it to be applicable to many other model organisms and most cells in tissue types that are well-vascularized that can be processed to single cells.
2. The reagents puromycin and digitonin vary somewhat from lot to lot and especially from the company purchased. We suggest piloting these reagents ahead of time in comparison experiments, and then ordering and freezing these reagents in bulk for future studies.
Acknowledgments
We thank Alexandre David for developing the original in vitro RPM protocol and Jaroslav Holly for helping with experiments.
Competing interests
The authors declare no conflict of interest.
Ethical considerations
All animal studies were approved by and performed in accordance with the Animal Care and Use Committee of the National Institute of Allergy and Infectious Diseases under protocol LVD-5E.
References
Hosios, A. M., Hecht, V. C., Danai, L. V., Johnson, M. O., Rathmell, J. C., Steinhauser, M. L., Manalis, S. R. and Vander Heiden, M. G. (2016). Amino Acids Rather than Glucose Account for the Majority of Cell Mass in Proliferating Mammalian Cells. Dev Cell. 36(5): 540–549. https://doi.org/10.1016/j.devcel.2016.02.012.
Mourant, J. R., Short, K. W., Carpenter, S., Kunapareddy, N., Coburn, L., Powers, T. M. and Freyer, J. P. (2005). Biochemical differences in tumorigenic and nontumorigenic cells measured by Raman and infrared spectroscopy. J Biomed Opt. 10(3): 031106. https://doi.org/10.1117/1.1928050.
Nathans, D. (1964). Puromycin Inhibition of Protein Synthesis: Incorporation of Puromycin into Peptide Chains. Proc Natl Acad Sci USA. 51(4): 585–592. https://doi.org/10.1073/pnas.51.4.585.
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. https://doi.org/10.1101/gad.219105.113.
David, A., Dolan, B. P., Hickman, H. D., Knowlton, J. J., Clavarino, G., Pierre, P., Bennink, J. R. and Yewdell, J. W. (2012). Nuclear translation visualized by ribosome-bound nascent chain puromycylation. J Cell Biol. 197(1): 45–57. https://doi.org/10.1083/jcb.201112145.
Seedhom, M. O., Hickman, H. D., Wei, J., David, A. and Yewdell, J. W. (2016). Protein Translation Activity: A New Measure of Host Immune Cell Activation. J Immun. 197(4): 1498–1506. https://doi.org/10.4049/jimmunol.1600088.
Graber, T. E., Hebert-Seropian, S., Khoutorsky, A., David, A., Yewdell, J. W., Lacaille, J. C. and Sossin, W. S. (2013). Reactivation of stalled polyribosomes in synaptic plasticity. Proc Natl Acad Sci USA. 110(40): 16205–16210. https://doi.org/10.1073/pnas.1307747110.
Seedhom, M. O., Dersh, D., Holly, J., Pavon-Eternod, M., Wei, J., Angel, M., Shores, L., David, A., Santos, J., Hickman, H., et al. (2024). Paradoxical imbalance between activated lymphocyte protein synthesis capacity and rapid division rate. eLife. 12. https://doi.org/10.7554/eLife.89015.
Aviner, R. (2020). The science of puromycin: From studies of ribosome function to applications in biotechnology. Comput Struct Biotechnol J. 18: 1074–1083. https://doi.org/10.1016/j.csbj.2020.04.014.
Arguello, R. J., Reverendo, M., Mendes, A., Camosseto, V., Torres, A. G., Ribas de Pouplana, L., van de Pavert, S. A., Gatti, E. and Pierre, P. (2018). SunRiSE - measuring translation elongation at single-cell resolution by means of flow cytometry. J Cell Sci. 131(10). https://doi.org/10.1242/jcs.214346.
Article Information
Publication history
Received: Sep 22, 2024
Accepted: Nov 17, 2024
Available online: Dec 12, 2024
Published: Jan 20, 2025
Copyright
© 2025 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
Cell Biology > Cell-based analysis > Flow cytometry
Cell Biology > Cell-based analysis > Organelle motility
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FlashTag-mediated Labeling for Intraventricular Macrophages in the Embryonic Brain
HA Hisa Asai
MO Mizuki Ono
TM Takaki Miyata
YH Yuki Hattori
Published: Vol 15, Iss 2, Jan 20, 2025
DOI: 10.21769/BioProtoc.5166 Views: 365
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Original Research Article:
The authors used this protocol in Cell Reports Feb 2023
Abstract
The fate mapping technique is essential for understanding how cells differentiate and organize into complex structures. Various methods are used in fate mapping, including dye injections, genetic labeling (e.g., Cre-lox recombination systems), and molecular markers to label cells and track their progeny. One such method, the FlashTag system, was originally developed to label neural progenitors. This technique involves injecting carboxyfluorescein diacetate succinimidyl ester (CFSE) into the lateral ventricles of mouse embryos, relying on the direct uptake of dye by cells. The injection of CFSE into the lateral ventricle allows for the pulse labeling of mitotic (M-phase) neural progenitors in the ventricular zone and their progeny throughout the brain. This approach enables us to trace the future locations and differentiation paths of neural progenitors. In our previous study, we adapted this method to selectively label central nervous system–associated macrophages (CAMs) in the lateral ventricle by using a lower concentration of CFSE compared to the original protocol. Microglia, the brain's immune cells, which play pivotal roles in both physiological and pathological contexts, begin colonizing the brain around embryonic day (E) 9.5 in mice, with their population expanding as development progresses. The modified FlashTag technique allowed us to trace the fate of intraventricular CAMs, revealing that certain populations of microglia are derived from these cells. The optimized approach offers deeper insights into the developmental trajectories of microglia. This protocol outlines the modified FlashTag method for labeling intraventricular CAMs, detailing the CFSE injection procedure, evaluation of CFSE dilution, and preparation of tissue for immunohistochemistry.
Key features
• This protocol builds upon the method developed by Govindan et al. and extends its application to intraventricular CAMs.
• This protocol allows for the cell fate tracking of intraventricular CAMs within 24 h.
• This protocol requires the technique of intraventricular injection of CFSE into embryonic brains.
Keywords: Microglia Central nervous system–associated macrophage Brain Macrophage Development Ventricle Cell fate Tracking Embryo Cortex
Graphical overview
FlashTag-mediated labeling for intraventricular macrophages
Background
The fate mapping technique is crucial for deciphering how cells differentiate and organize into complex structures. Various methods, such as dye injections, Cre-lox genetic labeling, and molecular markers, are used to track the progeny of cells. One of the earliest and most widely used techniques is BrdU (5-bromo-2'-deoxyuridine) birth dating, which labels newborn neural lineage cells at a specific time point [1,2]. BrdU is incorporated into cells during the S phase of DNA synthesis. When administered to pregnant dams either via intraperitoneal injection or drinking water, it leads to systemic labeling of all S-phase cells in the embryo. This results in no spatial restriction, meaning all cells born across all regions are labeled. Another technique, in utero electroporation, is used to label newborn neural progenitors. In this method, a plasmid encoding a reporter protein is injected into the lateral ventricle of an embryonic mouse, and electrical pulses are applied to facilitate its entry into the cells [3,4]. This method primarily targets M-phase cells but can also label S-phase cells, allowing for the labeling of a wide range of unsynchronized progenitors. However, the detection of labeled cells only occurs after the reporter protein is expressed, typically about 10 h later, making it challenging to study immediate events post-transduction [5].
To overcome the limitations of traditional birth-dating techniques, the FlashTag method was developed to track isochronic cohorts of ventricular zone (VZ)-born cells in the developing central nervous system (CNS) [6,7]. This method uses carboxyfluorescein diacetate succinimidyl ester (CFSE), a dye that fluoresces only after being processed within the cell. It allows for the specific labeling of M-phase neural progenitors lining the VZ, with intracellular fluorescence detectable within 20 min of injection. In the developing pallium, the labeling window is narrow, restricted to 1–2 h post-injection [7].
Notably, the FlashTag method can be optimized to label CNS-associated macrophages (CAMs), which reside in the ventricular space. Our previous research showed that intraventricular CAMs, which frequently infiltrate the developing pallium at embryonic day 12.5 (E12.5), differentiate into microglia after their infiltration in mice [8]. Microglia, the immune cells of the CNS, play essential roles throughout various life stages. For example, microglia shape neuronal circuits by pruning synapses and maintain homeostasis by clearing apoptotic cells and debris in the postnatal and adult brain, while contributing to neurogenesis by regulating differentiation and cell number [9-13]. Genetic fate mapping studies in mice have shown that microglia and CAMs share a common origin from erythromyeloid progenitors (EMPs), which are generated in the yolk sac between E7.5 and E8.5 and colonize the brain around E9.5 [14,15].
In our previous study, we optimized the FlashTag method by using a lower concentration of CFSE, enabling us to trace the fate of intraventricular CAMs. We discovered that certain microglia populations originate from these CAMs, which infiltrate the pallium around E12.5. This protocol is particularly useful for tracking the fate and behavior of intraventricular CAMs within a limited time window.
Materials and reagents
Biological materials
1. ICR mice (purchased from Japan SLC)
Reagents
1. CellTraceTM CFSE Cell Proliferation kit (for flow cytometry) (Thermo Fisher Scientific, catalog number: C34554)
2. Fast Green FCF (FUJIFILM Wako Pure Chemical Corp., catalog number: 061-00031)
3. Otsuka normal saline (Otsuka Pharmaceutical Factory, Inc., catalog number: 035-081517)
4. Sodium azide (FUJIFILM Wako Pure Chemical Corp., catalog number: 195-11091)
5. NaCl (sodium chloride) (FUJIFILM Wako Pure Chemical Corp., catalog number: 191-01665)
6. KCl (potassium chloride) (Sigma-Aldrich, catalog number: 24-4290-5)
7. Na2HPO4-12H2O (FUJIFILM Wako Pure Chemical Corp., catalog number: 196-02835)
8. NaH2PO4-2H2O (FUJIFILM Wako Pure Chemical Corp., catalog number: 192-02815)
9. KH2PO4 (FUJIFILM Wako Pure Chemical Corp., catalog number: 167-04241)
10. HCl (Sigma-Aldrich, catalog number: 13-1640-5)
11. NaOH (FUJIFILM Wako Pure Chemical Corp., catalog number: 194-18865)
12. Triton X-100 (Thermo Scientific, catalog number: A16046-0F)
13. Bovine serum albumin (Sigma-Aldrich, catalog number: A9418-50G)
14. Tissue-Tek O.C.T. compound (Sakura Finetek Japan Co., Ltd., catalog number: 4583)
15. Goat anti-CD206 pAb (1:300) (R&D systems, Minneapolis, catalog number: AF2535, RRID: AB_2063012)
16. Mouse anti-FITC mAb (1:400) (BioLegend, catalog number: 408301, RRID: AB_528900)
17. Rabbit anti-IBA1 pAb (1:1000) (FUJIFILM Wako Pure Chemical Corp., catalog number: 019-19741, RRID: AB_839504)
18. Rabbit anti-P2RY12 pAb (1:500) (AnaSpec, catalog number: 55043A, RRID: AB_2298886)
19. Donkey anti-Mouse IgG (H+L) highly cross-adsorbed secondary antibody, Alexa FluorTM 488 (1:1000) (Thermo Fisher Scientific, catalog number: A-21202, RRID: AB_141607)
20. Donkey anti-Rabbit IgG (H+L) highly cross-adsorbed secondary antibody, Alexa FluorTM 546 (1:1000) (Thermo Fisher Scientific, catalog number: A-10040, RRID: AB_2534016)
21. Donkey anti-Rabbit IgG (H+L) highly cross-adsorbed secondary antibody, Alexa FluorTM 647 (1:1000) (Thermo Fisher Scientific, catalog number: A-31573, RRID: AB_2536183)
22. Donkey anti-Goat IgG (H+L) cross-adsorbed secondary antibody, Alexa FluorTM 546 (1:1000) (Thermo Fisher Scientific, catalog number: A-11056, RRID: AB_2534103)
23. Donkey anti-Goat IgG (H+L) cross-adsorbed secondary antibody, Alexa FluorTM 647 (1:1000) (Thermo Fisher Scientific, catalog number: A-21447, RRID: AB_2535864)
24. Paraformaldehyde (Sigma-Aldrich, Merck, catalog number: 818715)
25. Sucrose (FUJIFILM Wako Pure Chemical Corp., catalog number: 190-00015)
26. Medetomidine hydrochloride (Nippon Zenyaku Kogyo Co., Ltd., catalog number: N.A.)
27. Midazolam (FUJIFILM Wako Pure Chemical Corp., catalog number: 135-13791)
28. Butorphanol tartrate (Meiji Animal Health Co., Ltd., catalog number: N.A.)
29. Atipamezole hydrochloride (Kyoritsu Seiyaku Corporation., catalog number: N.A.)
30. Fluoromount-G(R) (Cosmo Bio Co Ltd., catalog number: 0100-01)
31. 70% ethanol
Solutions
1. CFSE stock solution (see Recipes)
2. 0.3% Fast Green solution (see Recipes)
3. 10× PBS, pH 7.4 (see Recipes)
4. 1× PBS sterile, pH 7.4 (see Recipes)
5. 1× PBS with sodium azide, pH 7.4 (see Recipes)
6. Sodium azide solution (1% (w/v)) (see Recipes)
7. 2.5% CFSE working solution (see Recipes)
8. 5% CFSE working solution (see Recipes)
9. 10% CFSE working solution (see Recipes)
10. Three types of mixed anesthetic agents (see Recipes)
11. Anesthetic antagonist solution (see Recipes)
12. 0.2M phosphate buffer, pH 7.4 (see Recipes)
13. 8% PFA solution, pH 7.4 (see Recipes)
14. 4% PFA solution, pH 7.4 (see Recipes)
15. 20% sucrose solution (see Recipes)
16. Antibody diluent solution (see Recipes)
Recipes
1. CFSE stock solution (8 μL)
Reagent Final concentration Quantity or Volume
CellTraceTM CFSE (#C34554A) 6.25 mg/mL 50 μg
DMSO (attached in the kit; thaw if frozen) n/a 8 μL
Total n/a 8 μL
CFSE stock solution should be stored at -30 °C.
2. 0.3% Fast Green solution (100 mL)
Reagent Final concentration Quantity or Volume
Fast Green FCF 0.3% (w/v) 0.03 g
H2O (MilliQ water) n/a 10 mL
Total n/a 10 mL
0.3% Fast Green solution should be filtered through Millex-GV 0.22 μm filter and then stored in a 1.5 mL tube at 4 °C.
3. 10× PBS, pH 7.4 (2 L)
Reagent Final concentration Quantity or Volume
NaCl n/a 160 g
KCl n/a 4 g
Na2HPO4-12H2O n/a 58 g
KH2PO4 n/a 4 g
H2O (MilliQ water) n/a see note*
Total n/a 2 L
*Note: Allow the solution to mix completely and adjust the pH of the solution to 7.4 by adding 1M HCl and 1M NaOH while measuring with a pH meter. Then, bring the volume up to 2 L with MilliQ water.
4. 1× PBS sterile, pH 7.4 (500 mL)
Reagent Final concentration Quantity or Volume
10× PBS, pH 7.4 (Recipe 3) n/a 50 mL
H2O (MilliQ water) n/a 450 mL
Total n/a 500 mL see note*
*Note: Confirm that the pH of the solution is 7.4 using a pH meter. If it deviates, adjust it by adding 1M HCl or 1M NaOH. Then, bring the volume up to 500 mL with MilliQ water. The solution should be autoclaved for 20 min at 121 °C.
5. 1× PBS with sodium azide, pH 7.4 (500 mL)
Reagent Final concentration Quantity or Volume
1× PBS sterile, pH 7.4 (Recipe 4) n/a 500 mL
Sodium azide solution (1% w/v) (Recipe 6) 0.002% (w/v) 1 mL
Total n/a 501 mL
6. Sodium azide solution (1% w/v) (100 mL)
Reagent Final concentration Quantity or Volume
Sodium azide 1 g n/a
H2O (MilliQ water) n/a 100 mL
Total n/a n/a
The prepared solution should be stored at room temperature (RT).
7. 2.5% CFSE working solution (10 μL)
Reagent Final concentration Quantity or Volume
CFSE stock solution (Recipe 1) 2.5% (w/v) (156 μg/mL) 0.25 μL
1× PBS sterile, pH 7.4 (Recipe 4) n/a 8.75 μL
Fast Green solution (0.3% w/v) (Recipe 2) 0.03% (w/v) 1 μL
Total n/a 10 μL
8. 5% CFSE working solution (10 μL)
Reagent Final concentration Quantity or Volume
CFSE stock solution (Recipe 1) 5% (w/v) (313 μg/mL) 0.5 μL
1× PBS sterile, pH 7.4 (Recipe 4) n/a 8.5 μL
Fast Green solution (0.3% w/v) (Recipe 2) 0.03% (w/v) 1 μL
Total n/a 10 µL
9. 10% CFSE working solution (10 μL)
Reagent Final concentration Quantity or Volume
CFSE stock solution (Recipe 1) 10% (w/v) (625 µg/ml) 1 μL
1× PBS sterile, pH 7.4 (Recipe 4) n/a 8 μL
Fast Green Solution (0.3% w/v)
(Recipe 2)
0.03% (w/v) 1 μL
Total n/a 10 µL
10. Three types of mixed anesthetic agents (35 mL)
Reagent Final concentration Quantity or Volume
Midazolam n/a 40 mg
HCl (0.1M) n/a 2 mL see note*
Medetomidine hydrochloride
(1.0 mg/mL)
n/a 3 mL
Butorphanol tartrate (5.0 mg/mL) n/a 10 mL
Otsuka normal saline n/a 20 mL
Total n/a 35 mL
*Note: Midazolam needs to be diluted first using HCI (0.1M). After that, other reagents should be added.
Aliquot the prepared solution into 2 mL portions and store at 4 °C. Only the tube being used is stored at RT.
11. Anesthetic antagonist solution (20 mL)
Reagent Final concentration Quantity or Volume
Atipamezole hydrochloride
(5.0 mg/mL)
n/a 0.3 mL
Otsuka normal saline n/a 19.7 mL
Total n/a 20 mL
The prepared solution is aliquoted into 2 ml portions and stored at 4 °C. Only the tube being used is stored at RT.
12. 0.2 M phosphate buffer, pH 7.4 (1 L)
Reagent Final concentration Quantity or Volume
Na2HPO4-12H2O n/a 58 g
NaH2PO4-2H2O n/a 5.9 g
H2O (MilliQ water) n/a see note*
Total n/a 1 L
*Note: Allow the solution to mix completely and adjust the pH of the solution to 7.4 by adding 1M HCl and 1M NaOH while measuring with a pH meter. Then, bring the volume up to 1 L with MilliQ water.
13. 8% PFA solution, pH 7.4 (400 mL)
Reagent Final concentration Quantity or Volume
Paraformaldehyde 8% (w/v) 32 g
H2O (MilliQ water) n/a 365 mL see note*
10N NaOH n/a 90 μL see note**
H2O (MilliQ water) n/a see note**
Total n/a 400 mL
*Note: Add 32 g of paraformaldehyde to 365 mL of MilliQ water and dissolve it by heating it with a hot stirrer.
**Note: Add 90 μL of 10N NaOH at a temperature higher than 60 °C. When the cloudiness disappears and the solution becomes transparent, MilliQ water should be added. Confirm that the pH of the solution is 7.4 using a pH meter. If it deviates, adjust it by adding 1M HCl or 1M NaOH. Then, bring the volume up to 400 mL with MilliQ water and filter the solution with filter paper.
The prepared solution should be stored at 4 °C.
14. 4% PFA solution, pH 7.4 (200 mL)
Reagent Final concentration Quantity or Volume
8% PFA solution, pH 7.4 (Recipe 13) 4% (w/v) 100 mL
0.2M phosphate buffer, pH 7.4 (Recipe 12) n/a 100 mL
Total n/a 200 mL
The prepared solution should be stored at 4 °C.
15. 20% sucrose solution (250 mL)
Reagent Final concentration Quantity or Volume
1×PBS with sodium azide, pH 7.4 (Recipe 5) n/a 220 mL
Sucrose 20% 50 g
Total n/a 250 mL
The prepared solution should be stored at 4 °C.
16. Antibody diluent solution (100 mL)
Reagent Final concentration Quantity or Volume
1× PBS sterile, pH 7.4 (Recipe 4) n/a 100 mL
Triton X-100 0.1% (v/v) 100 μL
Sodium azide 0.1% (w/v) 0.1 g
Bovine serum albumin 3 mg/mL 0.3 g
Total n/a n/a
Filter the solution using Millex-HV 0.45 μm filter and store at 4 °C.
Laboratory supplies
1. Glass capillary with filament (NARISHIGE Group, catalog number: GD-1)
2. Eppendorf microloader (Merck, catalog number: EP5242956003)
3. Aspirator tube assembly (Drummondo, catalog number: 2-040-000)
4. Dropper silicone rubber for 2 mL (AZ ONE, catalog number: 6-356-02)
5. Heating pad (Koizumi, catalog number: N.A.)
6. Flat-bottom micro tube 1.5 mL large box of 500 × 20 bags (BIO-BIK, catalog number: CF-0150)
7. Corning 100 mm non-treated culture dish (Corning, catalog number: 430591)
8. Transfer pipette 2.3 mL (BM Equipment Co., Ltd., catalog number: 262-20S)
9. Extra fine Graefe forceps (Fine Science Tools, catalog number: 11151-10)
10. Fine Iris scissors (Fine Science Tools, catalog number: 14094-11)
11. Dumont #55 fine forceps (Fine Science Tools, catalog number: 11255-20)
12. Paper filter (As ONE, catalog number: 65-0426-67)
13. Millex-GV 0.22 μm filter (Millipore, catalog number: SLGVJ13SL)
14. Millex-HV 0.45 μm filter (Millipore, catalog number: SLHVR33RS)
15. Nipro Flomax hypodermic 26G (S.B) needle (Nipro, catalog number: 01046)
16. Terumo 1 mL syringe (Terumo, catalog number: SS-01T)
17. Nylon suture needles with thread (BEAR Medic Corporation, catalog number: SP15A05H-45)
18. MAS coat slide glass (Matsunami Glass Ind., Ltd., catalog number: SMAS-01)
19. NEO micro cover glass (Matsunami Glass Ind., Ltd., catalog number: C024501)
Equipment
1. Magnetic glass microelectrode horizontal puller (Narishige, model: PN-30)
2. Micro grinder (Narishige, model: EG-400)
3. e-HeatingBucket (Miracle beads bath, TAITEC Corp, model: BMB-17)
4. Confocal microscopy (Nikon, model: AXR)
5. Confocal microscopy (Nikon, model: TiE-A1R)
6. Cryostat (Leica Biosystems, model: CM1520)
Procedure
A. Fabrication of glass capillaries
1. Set the glass capillary in the magnetic glass microelectrode horizontal puller and pull it while heating to create two sharp-tipped capillaries from a single one (Figure 1A, B).
2. Break the tip of the glass capillary using forceps (Dumont #55 fine forceps) (Figure 1C, D).
3. Set the glass capillary in the micro grinder and adjust it to an angle of approximately 35° (Figure 1E).
4. Polish the tip of the glass capillary by touching it to the grinder surface several times (Figure 1F, G; Video 1).
Video 1. Polishing of a glass capillary. The video demonstrates the process of polishing the tip of a glass capillary using the micro grinder. The angle is adjusted to approximately 35°.
Figure 1. How to prepare glass capillaries and homemade pipette. A. The magnetic glass microelectrode horizontal puller is used to heat and pull a single glass capillary, creating two sharp-tipped capillaries. B. The glass capillary immediately after being processed with the puller. C. Picture showing how to cut the tip of the glass capillary using forceps. D. The glass capillary after the tip has been cut. The bottom image shows a magnified view of the cut tip. E. A glass capillary set in the micro grinder, with the angle adjusted to 35°. F. Polishing the tip of the glass capillary using the micro grinder. This process can be seen in Video 1. G. A magnified view of the completed glass capillary after all processing steps. Scale bar, 100 μm. H. Pictures demonstrating the process of creating the homemade pipette. The part of the aspirator tube assembly (indicated by the cyan circle in the left image) was removed and then inserted into the dropper silicone rubber for 2 mL (shown in the middle and right images).
B. Assembling the homemade pipette
1. Remove the part shown within the cyan circle in the left photo in Figure 1H from the aspirator tube assembly (Figure 1H).
2. Insert the removed part into the dropper silicone rubber for 2 mL (see the middle and right photos in Figure 1H).
C. Stock reagent preparation
1. Prepare the CFSE stock solution (see Recipe 1).
2. Prepare 0.3% Fast Green solution (See Recipe 2).
D. Preparation of CFSE working solutions
1. For labeling intraventricular CAMs, prepare the 2.5% CFSE working solution in a 1.5 mL tube and mix by pipetting (total 10 μL) (see Recipe 7) (Figure 2A, B). The solution should be prepared just before use.
2. Mark the glass capillaries. To measure 1 μL, mark 3.5 mm from the thickest part of the glass capillary, away from the tip, with an inner diameter of 0.6 mm (3.5 mm = 1 μL; 0.3 mm × 0.3 mm × 3.14 × 3.5 mm = 1.0 μL) (Figure 2C).
3. Using a microloader pipette, fill the marked glass capillaries with the 2.5% CFSE working solution (Figure 2D).
4. Attach the glass capillary filled with solution to the homemade pipette (Figure 2E).
Note: In this protocol paper, we prepared not only 2.5% but also 5% and 10% CFSE working solutions to examine concentration conditions to specifically label intraventricular macrophages. We found that 2.5% CFSE working solution can specifically label intraventricular macrophages in the left hemisphere [8]. The 10% solution matches the condition used in Govindan et al. [7].
E. Preparation for surgery
1. Warm saline to 37–40 °C in the beads bath.
2. Prepare surgical instruments (scissors, forceps).
3. Prepare three types of mixed anesthetic agents (see Recipe 10) and anesthetic antagonist solution (see Recipe 11).
F. Injection of the CFSE working solution
1. Anesthetize a pregnant ICR mouse at gestational day 12. Administer 80 μL of three types of mixed anesthetic agents (see Recipe 10) intraperitoneally using a 26G needle and a 1 mL syringe.
2. Wait 10 min for stable anesthesia.
3. Place the animal on the animal heating pad. After disinfecting the abdomen with 70% ethanol, dissect the abdomen of the dam using scissors, cutting the skin and then the peritoneum.
4. Extract the uterus and check the number and condition of the embryos.
5. Return the non-injected side of the uterus into the abdomen, exposing only the side for treatment (Figure 2F). Prevent drying by applying warmed saline as needed.
6. Confirm the fetal head orientation. Position the glass capillary filled with the CFSE working solution to pierce through the uterine wall, fetal skin, and into the brain ventricles.
7. After piercing the fetal brain, inject 1 μL of the CFSE working solution into the right lateral ventricle of each embryo using a glass capillary with the homemade pipette, guided by the 1 µL marking as a reference. Slight leakage into the left ventricle is acceptable (Figure 2G–I; Video 2).
8. Note the position of the treated embryo in the uterus in the log.
9. After injection, return the uterus to the abdomen and keep moist with saline.
10. If necessary, exteriorize the other uterus and perform the same procedure according to steps F4–9.
Video 2. Injection of the solution into the fetal mouse brain ventricles. This video shows the process of inserting the tip of a glass capillary filled with carboxyfluorescein diacetate succinimidyl ester (CFSE) working solution into the right brain of a fetal mouse through the maternal uterus and injecting the solution into the lateral ventricle.
Figure 2. Injection of carboxyfluorescein diacetate succinimidyl ester (CFSE) working solution into the lateral ventricle of the embryonic mouse. A. The CFSE stock solution was prepared by dissolving the CFSE provided in the kit using the attached DMSO. B. The 2.5% CFSE working solution was prepared in a 1.5 mL tube. C. Glass capillaries marked every 3.5 mm to measure out 1 μL of CFSE working solution. D. The CFSE working solution was loaded into the glass capillary using a microloader pipette. E. The glass capillary filled with the solution was attached to the homemade pipette holder. F. After anesthetizing the mother, one side of the uterus was exposed. G. 1 μL of the CFSE working solution was injected into the right lateral ventricle of the embryo located inside the uterus. H. An image taken immediately after the solution was injected into the right lateral ventricle of the embryo. The space in the right lateral ventricle was visualized by the green color of the Fast Green. A small amount of the solution leaked into the left lateral ventricle. I. A magnified image of the fetal head shown in H.
G. Suturing and recovery from anesthesia
1. Suture the peritoneum first and then the skin using Nylon suture needles with thread.
2. Administer 400 μL of anesthetic antagonist solution (see Recipe 11) to the dam intraperitoneally and place the mother mouse in a cage on a heated pad to recover.
H. Brain fixation
1. Two to twenty-four hours after injection, perform cervical dislocation of the mother mouse. Confirm maternal death, open the uterus and amniotic membrane, collect fetal heads or brains using forceps (Dumont #55 fine forceps), and rinse with 1× PBS sterile, pH 7.4.
2. Fix the fetal heads or brains in 4% PFA solution, pH 7.4 (see Recipe 14). For brains, immerse them in 4% PFA solution on ice for 1–1.5 h. For heads, immerse them in 4% PFA solution on ice for 2–3 h.
3. Transfer to 20% sucrose solution (see Recipe 15) on ice. Leave until the brain sinks (approximately 2 h).
4. Rinse lightly with O.C.T. compound to remove 20% sucrose solution.
5. Embed in O.C.T. compound in an appropriate mold, orient the brain under a microscope, and rapidly freeze with liquid nitrogen.
6. Store the embedded brain at -30 °C.
I. Cryosectioning, immunostaining, and observation
1. Prepare 16 μm cryosections of fetal brain tissue onto the slide glass (MAS coat slide glass) using a cryostat.
2. Dry sections as needed (store unused sections at -30 °C).
3. Incubate slides with cryosections in 1×PBS with sodium azide, pH 7.4 (see Recipe 5) to remove the O.C.T. compound (RT, 10 min).
4. Incubate with primary antibodies (4 °C, overnight).
5. Wash slides with 1× PBS with sodium azide, pH 7.4 (RT, 10 min, three times).
6. Incubate with secondary antibodies (RT, 1 h).
7. Wash slides with 1× PBS with sodium azide, pH 7.4 (RT, 10 min, three times).
8. Mount with mounting media [Fluoromount-G(R)] and coverslips (NEO micro cover glass).
9. Observe using confocal microscopy.
10. Evaluation of cell fate should be performed on the left hemisphere, where only intraventricular CAMs are labeled with CFSE (Figure 3, 4).
Note: For the detection of CFSE, the signal was enhanced by using mouse anti-FITC mAb as the primary antibody and donkey anti-mouse IgG (H+L) highly cross-adsorbed secondary antibody, Alexa FluorTM 488 as the secondary antibody.
Figure 3. Optimization of concentration conditions for carboxyfluorescein diacetate succinimidyl ester (CFSE) working solution. A. Image illustrating the experimental procedure for the FlashTag-based cell fate analysis for intraventricular central nervous system–associated macrophages (CAMs) for the E12.5 ICR mice. B. Immunostaining for FITC (to detect CFSE) and Iba1 (a marker for CAMs and microglia) in the brain of embryos 2 h after injecting the CFSE working solution into the right lateral ventricles. White dotted line, ventricular surface contour of the pallium. Scale bar, 100 μm.
Figure 4. FlashTag-based cell fate trace analysis of infiltrated intraventricular central nervous system–associated macrophages (CAMs) into microglia (DOI: 10.1016/j.celrep.2023.112092) A. Triple-fluorescence picture [CFSE, Iba1, and CD206 (a marker for CAMs)] of an E12.5 brain that was fixed 3 h after intraventricular injection with carboxyfluorescein diacetate succinimidyl ester (CFSE). The intraventricular CD206+Iba1+ cells on the inner surface of the pallium (yellow arrowhead) were CFSE+, whereas the pallial Iba1+CD206– cells (preexisting microglia) were negative for CFSE. B. Graph showing the proportion of the CFSE+Iba1+ cells per total Iba1+ cells comparing the ventricle and the pallium (two-sided Mann–Whitney U test; N = six male and female mice; the average value of six sections from each animal is plotted; P = 0.002). C–H. FlashTag-based analysis of the transition from the CAMs toward microglia, with immunofluorescence for CFSE and Iba1 (C) or P2RY12 (a marker for microglia) and CD206 (D) 2 h and 24 h after injection. Graphs showing the proportions of CFSE+Iba1+ cells among Iba1+ cells (E) and the CFSE+ cells that were also P2RY12+ (F), CD206+ (G), or CD206+P2RY12+ (H) [two-sided Steel–Dwass test; N = six male and female mice; the average value of six sections from each animal is plotted; P = 0.032, 0.032 in E, 0.031, 0.032 in F, 0.027, 0.031 in G, and 0.032, 0.051 in H (left-to-right)]. White broken line, ventricular surface contour of the pallium. Scale bar, 100 μm.
Data analysis
Quantitative data are presented as the mean value ± S.D. of representative experiments. Statistical differences between groups were analyzed using R software by the Mann–Whitney U test for two-group comparisons or the Steel–Dwass test for multiple comparisons. All the statistical tests were two-tailed, and P < 0.05 was considered to indicate statistical significance. The P value is shown in each graph (n.s., not significant). Individual values are plotted as circles in the bar graphs. The number of samples examined in each analysis is shown in the corresponding figure legend. No randomization was used, and no samples were excluded from the analysis. No statistical methods were used to predetermine the sample size owing to experimental limitations.
Validation of protocol
The protocol has been validated using multiple biological replicates (e.g., N = six mice per condition) to ensure reproducibility. Statistical analysis was performed using the Mann–Whitney U test for two-group comparisons or the Steel–Dwass test for multiple comparisons. The results confirm the reliability of this protocol and are detailed in the Data analysis section and the corresponding figure legend (refer to Figure 4).
This protocol or parts of it has been used and validated in the following research article:
Hattori et al. [8] CD206(+) macrophages transventricularly infiltrate the early embryonic cerebral wall to differentiate into microglia. Cell Reports (Figure 4, panel A–I)
General notes and troubleshooting
General notes
FlashTag was originally developed to trace the fate of neural progenitors. We modified this method by diluting the CFSE working solution to about one-fourth of its original concentration (2.5%), allowing us to specifically label intraventricular CAMs in the embryonic mouse brain. Since the injection is targeted into the right ventricle, the dye is taken up by not only the intraventricular CAMs but also the neural progenitors in the pallium on the right hemisphere. However, on the opposite side (the left, non-injected side), the dye selectively stains intraventricular CAMs, likely due to their higher phagocytic activity compared to other cells. We confirmed that almost all CAMs present in the left lateral ventricle were labeled with CFSE (Figure 4B). Similar staining has also been successfully performed at other stages (e.g., E13.5, E14.5). Since the infiltration of intraventricular CAMs frequently occurs at E12.5 in mice, we believe that labeling intraventricular CAMs at E12.5 is crucial for conducting fate mapping. By leveraging this feature, we successfully traced the fate of the intraventricular CAMs. However, as the dye is metabolized relatively quickly, tracking beyond 24 h appears challenging, though it is effective for tracing within that timeframe.
Troubleshooting
Problem 1: The CFSE working solution leaks out of the ventricle in the fetal mouse brain.
Possible cause(s): The tip of the glass capillary might not be inserted at the correct position. It could either be too shallow, causing the solution to enter subcutaneously, or inserted too deeply, resulting in injection outside the ventricle.
Solution(s): If the solution is properly injected into the ventricle, it will accumulate in the shape of the ventricular space, so confirming this clarity is an indicator of accuracy.
Problem 2: Even when using the 2.5% CFSE working solution, cells other than intraventricular CAMs are being labeled.
Possible cause(s): There may be variability in the amount of CFSE working solution administered.
Solution(s): Since our analysis is based on the left hemisphere at this concentration, please confirm that you are analyzing the left hemisphere. In the right hemisphere, other cells are also labeled. Additionally, there may be variations in the solution volume due to the tools used, so ensure accurate measurement. It may also be necessary to optimize the injection volume.
Problem 3: The embryos are dying.
Possible cause(s): The surgical procedure may be causing excessive stress.
Solution(s): Perform the procedure quickly to minimize the burden on the mouse. Experienced individuals can complete the procedure, from the incision to the final suturing, in approximately 20–30 min. Additionally, it is also necessary to handle the embryos with care.
Acknowledgments
We thank Makoto Masaoka, Namiko Noguchi, and Ikuko Mizuno (Department of Anatomy and Cell Biology, Nagoya University Graduate School of Medicine) for their technical assistance. We wish to acknowledge the Division for Medical Research Engineering, Nagoya University Graduate School of Medicine, for technical support. This work was supported by JSPS Grants-in-Aid for Scientific Research (B) [JP21H02656 (T.M.), JP23H02658 (Y.H.)], Grants-in-Aid for Transformative Research Areas (A) [JP23H04161 (Y.H.)], and JST FOREST [JPMJFR214C (Y.H.)]. This study was also supported by grants from The Uehara Memorial Foundation, Takeda Science Foundation, Inoue Foundation for Science, Tokai Pathways to Global Excellence (T-GEx), and AMED-ASPIRE program [JP23jf0126004 (Y.H.)]. This protocol was described and validated in Hattori et al., Cell Reports (2023) (DOI: 10.1016/j.celrep.2023.112092).
Competing interests
The authors declare no competing interests.
Ethical considerations
1. The animal experiments were conducted according to the Japanese Act on Welfare and Management of Animals, Guidelines for Proper Conduct of Animal Experiments (published by the Science Council of Japan), and Fundamental Guidelines for Proper Conduct of Animal Experiment and Related Activities in Academic Research Institutions (published by the Ministry of Education, Culture, Sports, Science and Technology, Japan).
2. All protocols for animal experiments were approved by the Institutional Animal Care and Use Committee of Nagoya University (No. 29006).
References
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Rochefort, C., Gheusi, G., Vincent, J. D. and Lledo, P. M. (2002). Enriched odor exposure increases the number of newborn neurons in the adult olfactory bulb and improves odor memory. J Neurosci. 22(7): 2679-2689. https://doi.org/10.1523/JNEUROSCI.22-07-02679.2002.
Tabata, H. and Nakajima, K. (2001). Efficient in utero gene transfer system to the developing mouse brain using electroporation: visualization of neuronal migration in the developing cortex. Neuroscience. 103(4): 865-872. https://doi.org/10.1016/s0306-4522(01)00016-1.
LoTurco, J., Manent, J. B. and Sidiqi, F. (2009). New and improved tools for in utero electroporation studies of developing cerebral cortex. Cereb Cortex. 19 Suppl 1(Suppl 1): i120-125. https://doi.org/10.1093/cercor/bhp033.
Stancik, E. K., Navarro-Quiroga, I., Sellke, R. and Haydar, T. F. (2010). Heterogeneity in ventricular zone neural precursors contributes to neuronal fate diversity in the postnatal neocortex. J Neurosci. 30(20): 7028-7036. https://doi.org/10.1523/JNEUROSCI.6131-09.2010.
Telley, L., Govindan, S., Prados, J., Stevant, I., Nef, S., Dermitzakis, E., Dayer, A. and Jabaudon, D. (2016). Sequential transcriptional waves direct the differentiation of newborn neurons in the mouse neocortex. Science 351(6280): 1443-1446. https://doi.org/10.1126/science.aad8361.
Govindan, S., Oberst, P. and Jabaudon, D. (2018). In vivo pulse labeling of isochronic cohorts of cells in the central nervous system using FlashTag. Nat Protoc. 13(10): 2297-2311. https://doi.org/10.1038/s41596-018-0038-1.
Hattori, Y., Kato, D., Murayama, F., Koike, S., Asai, H., Yamasaki, A., Naito, Y., Kawaguchi, A., Konishi, H., Prinz, M., et al. (2023). CD206(+) macrophages transventricularly infiltrate the early embryonic cerebral wall to differentiate into microglia. Cell Rep. 42(2): 112092. https://doi.org/10.1016/j.celrep.2023.112092.
Li, Q. and Barres, B. A. (2018). Microglia and macrophages in brain homeostasis and disease. Nat Rev Immunol 18(4): 225-242. https://doi.org/10.1038/nri.2017.125.
Paolicelli, R. C., Bolasco, G., Pagani, F., Maggi, L., Scianni, M., Panzanelli, P., Giustetto, M., Ferreira, T. A., Guiducci, E., Dumas, L., et al. (2011). Synaptic pruning by microglia is necessary for normal brain development. Science. 333(6048): 1456-1458. https://doi.org/10.1126/science.1202529.
Hattori, Y. (2021). The behavior and functions of embryonic microglia. Anat Sci Int. https://doi.org/10.1007/s12565-021-00631-w.
Hattori, Y. and Miyata, T. (2018). Microglia extensively survey the developing cortex via the CXCL12/CXCR4 system to help neural progenitors to acquire differentiated properties. Genes Cells. 23(10): 915-922. https://doi.org/10.1111/gtc.12632.
Parkhurst, C. N., Yang, G., Ninan, I., Savas, J. N., Yates, J. R., 3rd, Lafaille, J. J., Hempstead, B. L., Littman, D. R. and Gan, W. B. (2013). Microglia promote learning-dependent synapse formation through brain-derived neurotrophic factor. Cell. 155(7): 1596-1609. https://doi.org/10.1016/j.cell.2013.11.030.
Ginhoux, F., Greter, M., Leboeuf, M., Nandi, S., See, P., Gokhan, S., Mehler, M. F., Conway, S. J., Ng, L. G., Stanley, E. R., et al. (2010). Fate mapping analysis reveals that adult microglia derive from primitive macrophages. Science. 330(6005): 841-845. https://doi.org/10.1126/science.1194637.
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Article Information
Publication history
Received: Sep 22, 2024
Accepted: Nov 28, 2024
Available online: Dec 12, 2024
Published: Jan 20, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
How to cite
Category
Neuroscience > Development > Neuron
Developmental Biology > Cell growth and fate > Differentiation
Immunology > Immune cell function > Macrophage
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Cochlear Organ Dissection, Immunostaining, and Confocal Imaging in Mice
CC Chenyu Chen *
BC Binjun Chen *
XQ Xiaoqing Qian
HS Haojie Sun
XF Xiao Fu
DR Dongdong Ren
(*contributed equally to this work)
Published: Vol 15, Iss 2, Jan 20, 2025
DOI: 10.21769/BioProtoc.5167 Views: 1603
Reviewed by: Marion HoggMohammed Mostafizur Rahman Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in eNeuro Jun 2023
Abstract
The organ of Corti, located in the inner ear, is the primary organ responsible for animal hearing. Each hair cell has a V-shaped or U-shaped hair bundle composed of actin-filled stereocilia and a kinocilium supported by true transport microtubules. Damage to these structures due to noise exposure, drug toxicity, aging, or environmental factors can lead to hearing loss and other disorders. The challenge when examining auditory organs is their location within the bony labyrinth and their small and fragile nature. This protocol describes the dissection procedure for the cochlear organ, followed by confocal imaging of immunostained endogenous and fluorescent proteins. This approach can be used to understand hair cell physiology and the molecular mechanisms required for normal hearing.
Key features
• Protocol for the microdissection of the organ of Corti and suitable preparation for later immunostaining.
• This technique involves the evaluation of mouse cochlea for planar-cell-polarity protein.
• Quantitative and qualitative analysis of hair cell cilia in different dimensions.
Keywords: Cochlea Hair cells Dissection Immunostaining Confocal Imaging
Graphical overview
Background
The inner ear, serving as both the auditory organ and local sensory receptor, houses the critical auditory sensor [1,2]. It consists of three rows of outer hair cells and one row of inner hair cells, as well as various supporting cells. These cellular components are meticulously arrayed in a mosaic pattern on the intricate cochlear membrane of the inner ear, demonstrating remarkable flexibility [3–6]. Hair cells convert acoustic signals into electrical signals through the deflection of tip cilia. These cilia are important cellular structures involved in normal morphogenesis and functional activities of hair cells [7,8]. Electrical impulses are ultimately channeled to the cerebral cortex, facilitating auditory perception. Research shows that the primary cause of hearing loss is the involuntary regeneration of damaged hair cells in the mammalian inner ear [9]. The investigation into the auditory impairment mechanism hinges upon the intricate functioning of the Corti organ. Advances in employing specialized immunochemical techniques have significantly enhanced our comprehension of its fundamental operating principles (Figure 1).
Figure 1. Inner ear anatomy. A, B) The cochlea and a close-up of a single turn cross-section, revealing the organ of Corti. C) The organ of Corti contains mechanosensitive hair cells, which lie beneath the tectorial membrane. D) Hair bundles composed of actin-filled stereocilia arranged in a staircase configuration.
Hair cells exhibit a unique form of planar cell polarity, with their stereocilia bundles oriented toward the peripheral region of the cochlear duct, characterized by a distinct alignment [10]. In this process, several proteins have to be targeted to specific cellular locations to signal the directionality in the cells and to build the specific polarity structure that is unique for the maintenance of the mechanotransduction apparatuses of the sensory hair cells [11–13]. Immunohistochemical techniques are employed to discern hair cell types and precisely locate native proteins contributing to cellular architecture and mechanotransduction capabilities. Furthermore, confocal microscopy enables precise visualization of the ultrastructural features and properties of cochlear lesions, thereby facilitating comprehension of auditory physiological and pathological dynamics. This innovative experimental methodology is extensively employed in fundamental research on auditory sensory cells within the cochlea. We hereby propose a comprehensive protocol designed for the meticulous examination of the mouse auditory organ, encompassing its anatomical structure, immunohistochemical staining procedures, and advanced confocal imaging techniques.
Materials and reagents
Biological materials
1. Rab11a conditional knock-out alleles, Vangl2-Looptail mice (The Jackson Laboratory, catalog number: 000220)
Reagents
1. Phosphate-buffered saline (PBS) (GENOM, catalog number: GNM20012-5)
2. Paraformaldehyde (PFA) (Sigma-Aldrich, catalog number: 158127)
3. Triton X-100 (Sigma, catalog number: 9002-93-1)
4. Normal donkey serum (Merck, Millipore, catalog number: S30)
5. Primary antibodies:
a. Rab11a, 1:200 (Cell Signaling Technology, catalog number: 2413)
b. γ-tubulin, 1:200 (Sigma, catalog number: T6557)
c. Arl13b, 1:1500 (Tamara Caspary, Emory University, Atlanta, GA)
d. Vangl2, 1:200 (R&D Systems, catalog number: AF4815)
e. Fz3, 1:500 (gift from Jeremy Nathans, Johns Hopkins University, Baltimore, MD)
f. LGN, 1:200 (gift from Fumio Matsuzaki, RIKEN)
g. β-Spectrin, 1:200 (BD Transduction Laboratories, catalog number: 612562)
h. MyosinVIIa, 1:200 (Proteus Bioscience Inc, catalog number: 25-6790)
i. Radixin, 1:100 (Abcam, catalog number: ab52495)
j. E-Cadherin, 1:200 (Invitrogen, catalog number: 13-1700)
6. Secondary antibodies:
a. Alexa Fluor® 488 AffiniPure donkey anti-mouse IgG (H+L), 1:1,000 (Jackson Immuno-Research Laboratories, catalog number: 715-545-151)
b. Rhodamine RedTM-X AffiniPure donkey anti-mouse IgG (H+L), 1:1,000 (Jackson Immuno-Research Laboratories, catalog number:715-295-151)
c. Alexa Fluor® 647 AffiniPure donkey anti-mouse IgG (H+L), 1:1,000 (Jackson Immuno-Research Laboratories, catalog number:715-605-151)
7. Phalloidin, 1:1,000 (Sigma-Aldrich, catalog number: P5282)
8. Mounting medium (Vectashield Antifade Medium, catalog number: H-1200)
9. Sodium chloride (NaCl) (Sigma-Aldrich, catalog number: S3014)
10. Potassium chloride (KCl) (Sigma-Aldrich, catalog number: P9541)
11. Sodium phosphate dibasic heptahydrate (NaH2PO4·7H2O) (Sigma-Aldrich, catalog number: S9390)
12. Potassium dihydrogen phosphate (KH2PO4) (Sigma-Aldrich, catalog number: P9791)
Solutions
1. 1× PBS (see Recipes)
2. 4% PFA (see Recipes)
Recipes
1. 1× PBS
Reagent Final concentration Amount
Sodium chloride n/a 8 g
Potassium chloride n/a 200 mg
Sodium phosphate dibasic heptahydrate n/a 1.44 g
Potassium dihydrogen phosphate n/a 240 mg
H2O n/a 1,000 mL
Total n/a 1,000 mL
Adjust pH to 7.4 with HCl and store it at room temperature.
2. 4% PFA
Reagent Final concentration Amount
Paraformaldehyde (powder) n/a 4 g
1× PBS n/a 100 mL
Total 4% 100 mL
Store paraformaldehyde powder at 4 °C. Heat while stirring under the chemical hood at approximately 60 °C. When the solution is transparent, let it cool down and then filter with Whatman paper to remove undissolved particles. Freeze PFA aliquots in 50 mL tubes at -20 °C. Remember to wear suitable personal protective equipment.
Equipment
1. Dissection microscope (BT, catalog number: BTS-300)
2. Large forceps (Fine Science Tools, catalog number: 11026-15)
3. Fine forceps (Fine Science Tools, catalog number: 11252-00)
4. Very fine forceps (Fine Science Tools, catalog number: 11200-14)
5. Small scissors (Fine Science Tools, catalog number: 91460-11)
6. Confocal microscope (Carl Zeiss Microscopy, model: LSM 510, AxioObserver)
7. 48-well plate (Corning, catalog number: 3548)
8. Petri dish (BD Biosciences, Falcon®, catalog number: 351006)
Software and datasets
1. ImageJ software (http://rsb.info.nih.gov/ij)
Procedure
Below, we describe the step-by-step procedure for cochlear organ dissection, immunostaining, and confocal imaging in mice from E18.5 to P2.
A. Cochlear organ dissection
1. Euthanize the mouse via cervical dislocation or CO2 inhalation (100% CO2).
2. Decapitate the mouse with small scissors. Using large forceps, remove the brain on each side to identify the temporal bones.
3. Under a dissecting microscope, use fine forceps to isolate the bony labyrinths from the temporal bones.
4. Trim away additional tissue and remove the stapes from the oval window.
5. Use the tip of the fine forceps to clear the oval and round windows and make a small hole in the apex of the cochlear spiral. Fix the bony labyrinths in 1.5 mL of 4% PFA for 5 min on ice.
6. Place the bony labyrinths in a dissecting dish containing PBS to facilitate the next step (Figure 2A).
Figure 2. Cochlear organ dissection. A) After the removal of additional tissue from the temporal bone, the otic vesicles are revealed under a light microscope. B) Dissected organ of Corti.
7. Use fine forceps to remove the outer cartilage to expose the cochlear duct.
8. Use fine forceps to slowly remove the stria vascularis. Remove the whole stria in one attempt as far as possible.
9. Use very fine forceps to remove the Reissner’s membrane. Visualize the dorsal aspect of the cochlear duct, including the sensory epithelium.
10. Use very fine forceps to remove the tectorial membrane.
11. Use very fine forceps to remove the intact organ of Corti (Figure 2B, Video 1).
Video 1. Fine dissection of the organ of Corti
12. Fix the organ of Corti immediately in the 48-well plate containing 0.5 mL of 4% PFA for 2 h at room temperature (RT), 2 h on ice, or overnight at 4 °C.
B. Immunostaining
1. Transfer the organ of Corti to a different well in the 48-well plate containing 0.5 mL of 0.1% Triton X-100 in PBS (PBS-T) for 10–20 min at RT. Agitation is not required for any of these steps. Transfer the organ of Corti with very fine forceps.
2. Place the tissues in 10% normal donkey serum and PBS-T for 1 h at RT.
3. Incubate the organ of Corti tissues in primary antibody in 5% normal donkey serum and PBS-T overnight at 4 °C.
4. Wash with 0.5 mL of PBS-T three times, 1–2 h each, at RT.
5. Incubate the samples in secondary antibodies in 5% normal donkey serum and PBS-T for 2 h at RT or overnight at 4 °C. This procedure should be shielded from light.
6. Wash the sample with PBS-T three times every 1–2 h at RT.
7. Incubate the tissue in phalloidin in 5% normal donkey serum and PBS-T for 30 min at RT.
8. Wash the sample with PBS-T three times every 1–2 h at RT.
C. Confocal imaging
1. Gently transfer the basilar membrane into a drop of PBS placed on a microscope slide.
2. Under the microscope, use very fine forceps to orient the sample facing upward toward the coverslip.
3. Add a drop of mounting medium directly onto the basilar membrane sample. Place one side of the coverslip at an angle against the slide, making contact with the outer edge of the liquid drop, and then lower the cover gently. Take care to avoid air bubbles.
4. Use an adsorbent tissue or a piece of filter paper to remove any excess mounting medium from the edges of the coverslip.
5. Seal the coverslip to the glass slide with nail polish. The sample should be stored at 4 °C in a dark environment until observation.
6. Check the intensity of the labeling and evaluate the background of the experiment by looking at the negative controls.
7. Acquire all pictures in the same conditions (exposure and general settings) for each color channel.
8. Visualize with Zeiss LSM510 confocal microscope equipped with the ZEN software using 20–63× objectives (oil immersion, NA 1.4) and the 488- and 555-nm excitation wavelengths of the laser.
9. Use the Z projection function to provide a two-dimensional view of all the pictures of an image stack by projecting them along the axis perpendicular to the image plane. The Maximum Intensity option creates an image in which each pixel contains the maximum value over all images in the stack.
10. Define specific positions along the cochlear duct, such as 25%, 50%, and 75% from the base, and use these to compare the proteins of interest in cochlear regions.
11. Take pictures using the microscope software for quantification of the particular phenotype as required.
a. Stereocilia morphology: Measure the bundle convexity or the height, defined as the shortest distance between the vertex of the bundle and the line that connects two ends of the bundle. For each genotype, three independent cochleae were counted and quantified, noting the region and hair cell type.
b. Stereocilia orientation: Assess the orientation of each individual bundle relative to a line perpendicular to the planar axis. The angle is defined between the center line and the horizontal line. One line is drawn to bisect the middle of the V-shaped stereocilia bundle, and another line is drawn perpendicular to the planar axis.
c. Cilia quantification: Cilia marked by Arl13b immunostaining are recorded and counted according to different IHC rows and OHC rows. A minimum of 300 IHCs and 1,200 OHCs are counted per region of each genotype. Five individual animal samples are included in the quantification.
12. The above procedures are completed using ImageJ software. The specific steps of the software are as follows (Figure 3):
a. Open the images in ImageJ.
b. Use the straight line tool from the toolbar to connect two ends of the bundle.
c. Use the straight line tool from the toolbar to measure the shortest distance between the vertex of the bundle and the line.
d. Click Analyze > Measure and the length value will appear.
e. Use the angle tool from the toolbar to measure the V-shaped stereocilia bundle.
f. Use the straight line tool from the toolbar to bisect the middle of the V-shaped stereocilia bundle (angle 1 and angle 2 are equal).
g. Use the angle tool from the toolbar to measure the angle between the bisect line and the horizontal line (angle 3).
h. Click Analyze > Measure and the angle value will appear.
Figure 3. Cochlear hair cells and quantification of the particular aspects. A) Phalloidin staining (green) in the cochlea, showing the stereocilia bundles of hair cells. B) Zoomed-in view of a single hair cell. C) Schematic diagram of length measurement. The red line indicates the length. D) Schematic diagram of angle measurement. One line indicates the bisector of the angle in V-shaped stereocilia bundles (angle 1 and angle 2 are equal). The other line indicates the horizontal line (angle 3, marked in red, indicates the measurement angle).
Validation of protocol
This protocol or parts of it has been used and validated in the following research article:
Knapp et al. [14]. Rab11a Is Essential for the Development and Integrity of the Stereocilia and Kinocilia in the Mammalian Organ of Corti. eNeuro (Figure 1, panels A–D; Figure 4, panels A–E; Figure 5, panels A–H; Figure 6, panels A and B; Figure 7, panels F–I).
General notes and troubleshooting
Troubleshooting
Problem: Unsatisfactory immunostaining results.
Possible cause: Poor quality of samples collected; poor quality of antibodies used; irregular staining procedures.
Solution: Fresh samples should be used as much as possible to prevent problems such as degradation of proteins and severe dehydration. Ensure that the antibody is within its expiration date and use the same batch of antibody as much as possible. Note that it is necessary to avoid light when staining the secondary antibody.
Acknowledgments
Funding was provided by the National Natural Science Foundation of China (NSFC, Grants 82271166, 81970880, and 81771017 to D.R., 82201288 to X.Q.) and by the Natural Science Foundation of Shanghai (Grant No. 22ZR1410100 to D.R.). We thank eNeuro Journal for some illustrating images we used in this paper [14].
Competing interests
The authors declare no competing interests.
Ethical considerations
All procedures were approved by the Emory University Institutional Animal Care and Use committee and met the NIH guidelines.
References
Jang, M. W., Lim, J., Park, M. G., Lee, J. and Lee, C. J. (2022). Active role of glia‐like supporting cells in the organ of Corti: Membrane proteins and their roles in hearing. Glia. 70(10): 1799–1825.
Bieniussa, L., Jain, I., Bosch Grau, M., Juergens, L., Hagen, R., Janke, C. and Rak, K. (2023). Microtubule and auditory function – an underestimated connection. Semin Cell Dev Biol. 137: 74–86.
Xia, A., Udagawa, T., Quiñones, P. M., Atkinson, P. J., Applegate, B. E., Cheng, A. G. and Oghalai, J. S. (2022). The impact of targeted ablation of one row of outer hair cells and Deiters’ cells on cochlear amplification. J Neurophysiol. 128(5): 1365–1373.
Buswinka, C. J., Rosenberg, D. B., Simikyan, R. G., Osgood, R. T., Fernandez, K., Nitta, H., Hayashi, Y., Liberman, L. W., Nguyen, E., Yildiz, E., et al. (2024). Large-scale annotated dataset for cochlear hair cell detection and classification. Sci Data. 11(1): 416.
Prajapati-DiNubila, M., Benito-Gonzalez, A., Golden, E. J., Zhang, S. and Doetzlhofer, A. (2019). A counter gradient of Activin A and follistatin instructs the timing of hair cell differentiation in the murine cochlea. eLife. 8: e47613.
Wan, L., Lovett, M., Warchol, M. E. and Stone, J. S. (2020). Vascular endothelial growth factor is required for regeneration of auditory hair cells in the avian inner ear. Hear Res. 385: 107839.
Liu, Y., Qi, J., Chen, X., Tang, M., Chu, C., Zhu, W., Li, H., Tian, C., Yang, G., Zhong, C., et al. (2019). Critical role of spectrin in hearing development and deafness. Sci Adv. 5(4): eaav7803.
Dong, S. H., Kim, S. S., Kim, S. H. and Yeo, S. G. (2020). Expression of aquaporins in inner ear disease. Laryngoscope. 130(6): 1532–1539.
McQuate, A., Knecht, S. and Raible, D. W. (2023). Activity regulates a cell type-specific mitochondrial phenotype in zebrafish lateral line hair cells. eLife. 12: e80468.
Scheffer, D. I., Zhang, D. S., Shen, J., Indzhykulian, A., Karavitaki, K. D., Xu, Y. J., Wang, Q., Lin, J. C., Chen, Z. Y., Corey, D. P., et al. (2015). XIRP2, an Actin-Binding Protein Essential for Inner Ear Hair-Cell Stereocilia. Cell Rep. 10(11): 1811–1818.
Yu, D., Deng, D., Chen, B., Sun, H., Lyu, J., Zhao, Y., Chen, P., Wu, H. and Ren, D. (2022). Rack1 regulates cellular patterning and polarity in the mouse cochlea. Exp Cell Res. 421(2): 113387.
Okamoto, S., Chaya, T., Omori, Y., Kuwahara, R., Kubo, S., Sakaguchi, H. and Furukawa, T. (2017). Ick Ciliary Kinase Is Essential for Planar Cell Polarity Formation in Inner Ear Hair Cells and Hearing Function. J Neurosci. 37(8): 2073–2085.
Krey, J. F., Chatterjee, P., Dumont, R. A., O’Sullivan, M., Choi, D., Bird, J. E. and Barr-Gillespie, P. G. (2020). Mechanotransduction-Dependent Control of Stereocilia Dimensions and Row Identity in Inner Hair Cells. Curr Biol. 30(3): 442–454.e7.
Knapp, L., Sun, H., Wang, Y. M., Chen, B. J., Lin, X., Gao, N., Chen, P. and Ren, D. (2023). Rab11a Is Essential for the Development and Integrity of the Stereocilia and Kinocilia in the Mammalian Organ of Corti. eNeuro. 10(6): ENEURO.0420–22.2023.
Article Information
Publication history
Received: Sep 15, 2024
Accepted: Nov 28, 2024
Available online: Dec 10, 2024
Published: Jan 20, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
How to cite
Category
Cell Biology > Tissue analysis > Tissue isolation
Neuroscience > Basic technology > Tissue dissection
Cell Biology > Cell structure
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Using DIMet for Differential Analysis of Labeled Metabolomics Data: A Step-by-step Guide Showcasing the Glioblastoma Metabolism
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DOI: 10.21769/BioProtoc.5168 Views: 1001
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Original Research Article:
The authors used this protocol in Bioinformatics May 2024
Abstract
Stable-isotope resolved metabolomics (SIRM) is a powerful approach for characterizing metabolic states in cells and organisms. By incorporating isotopes, such as 13C, into substrates, researchers can trace reaction rates across specific metabolic pathways. Integrating metabolomics data with gene expression profiles further enriches the analysis, as we demonstrated in our prior study on glioblastoma metabolic symbiosis. However, the bioinformatics tools for analyzing tracer metabolomics data have been limited. In this protocol, we encourage the researchers to use SIRM and transcriptomics data and to perform the downstream analysis using our software tool DIMet. Indeed, DIMet is the first comprehensive tool designed for the differential analysis of tracer metabolomics data, alongside its integration with transcriptomics data. DIMet facilitates the analysis of stable-isotope labeling and metabolic abundances, offering a streamlined approach to infer metabolic changes without requiring complex flux analysis. Its pathway-based "metabologram" visualizations effectively integrate metabolomics and transcriptomics data, offering a versatile platform capable of analyzing corrected tracer datasets across diverse systems, organisms, and isotopes. We provide detailed steps for sample preparation and data analysis using DIMet through its intuitive, web-based Galaxy interface. To showcase DIMet's capabilities, we analyzed LDHA/B knockout glioblastoma cell lines compared to controls. Accessible to all researchers through Galaxy, DIMet is free, user-friendly, and open source, making it a valuable resource for advancing metabolic research.
Key features
• Glioblastoma tumor spheroids in vitro replicate tumors’ three-dimensional structure and natural nutrient, metabolite, and gas gradients, providing a more realistic model of tumor biology.
• Joint analysis of tracer metabolomics and transcriptomics datasets provides deeper insights into the metabolic states of cells.
• DIMet is a web-based tool for differential analysis and seamless integration of metabolomics and transcriptomics data, making it accessible and user-friendly.
• DIMet enables researchers to infer metabolic changes, offering intuitive and visually appealing "metabologram" outputs, surpassing conventional visual representations commonly used in the field.
Keywords: Metabolomics Glioblastoma Transcriptomics Differential analysis Data integration Bioinformatics
Graphical overview
Metabolomics and transcriptomics acquired data undergo downstream analysis with DIMet in Galaxy (image created with BioRender and Inkscape)
Background
Metabolic processes within cells play a major role in tumor adaptation, growth, and proliferation [1]. In the case of glioblastoma (GB), our previous in vitro studies focused on tumor plasticity driven by the hypoxic and acidic microenvironment and highlighted the critical role of lactate dehydrogenases (LDHA, LDHB) in this metabolic symbiosis [2]. We employed omics technologies—specifically transcriptomics and metabolomics—because they provide complementary insights into metabolic pathways. Transcriptomics captures the initial stage of gene expression through the analysis of mRNA transcripts, while metabolomics sheds light on the enzymatic activities associated with transcript translation.
To further characterize metabolic dynamics, we used stable-isotope resolved metabolomics (SIRM), also referred to as tracer or labeled metabolomics [3]. SIRM involves the administration of a stable isotope–labeled substrate to the cells or organisms, allowing to trace the integration of these isotopes into metabolic pathways, thus revealing subsequent biochemical reactions [3–5]. While 13C is the most commonly used isotope, other isotopes are also employed. The resulting corrected data contain the total abundances and labeling information, such as isotopologues and mean enrichment [4,6]. However, previous bioinformatics solutions for analyzing these datasets have exhibited limitations, including statistical frameworks or accessibility [7–11]. Furthermore, flux estimation often requires substantial computational resources and complex parameterization [12,13], creating a barrier to its adoption.
This protocol begins with the preparation of glioblastoma cell lines. Two distinct cell lines are used, P3 sgCont and sgLDHA/B, both generated from P3 wild-type glioblastoma cells through CRISPR-Cas9 editing [2]. For simplicity, we refer to sgCont as control throughout this paper. The sgLDHA/B cell line corresponds to a double knockout (KO) of the LDHA and LDHB genes and is referred to as LDHAB-KO. Along these cell lines, we provide the corresponding SIRM and transcriptomics datasets.
The primary focus of this paper is the data analysis using our recently published tool, DIMet [6], available in its Galaxy interface. DIMet is the first comprehensive tool designed for the differential analysis of tracer metabolomics data integrated with transcriptomics data. This protocol provides a detailed step-by-step guide for using DIMet in Galaxy. Specifically, we performed differential analyses between conditions and/or time points to evaluate total metabolite abundances and isotope labeling. By interpreting these two levels of information together, it becomes possible to infer variations in the production and consumption rates of metabolites, providing a proxy for flux differences [4,9].
We further combined metabolomics and transcriptomics datasets into DIMet's pathway-based metabolograms, addressing the limitations of previous tools. DIMet offers several key advantages: (i) it accommodates the unique requirements of SIRM data, (ii) it integrates data in a pathway-centric manner, (iii) it generates visually ergonomic outputs, and (iv) all functionalities are accessible via the free, user-friendly Galaxy interface. While complex analyses involving multiple isotopes and substrates are not yet supported, DIMet’s features and accessibility more than compensate for this limitation. Without restrictions on the system, organism, or isotope-substrate combination, DIMet in Galaxy can be used for differential analysis of any corrected tracer metabolomics dataset.
Materials and reagents
Biological materials
1. P3 glioblastoma stem-like cells (generated from patient [14] and modified through CRISPR-cas9 editing [2])
Reagents
1. Neurobasal medium (NBM) (Gibco, catalog number: 2110349), storage at 4 °C
2. B27 50× (Thermo Fisher Scientific, catalog number: 17504044), storage at 4 °C
3. FGF basic (PeproTech, catalog number: 100-18B), storage at -20 °C
4. Heparin (Sigma-Aldrich, catalog number: H3149)
5. Penicillin-streptomycin (Dutscher, catalog number: P06-07100), storage at -20 °C
6. Neurobasal medium without glucose (NBMW/O) (Gibco, catalog number: A2477501), storage at 4 °C
7. [13C]6-glucose (Sigma, catalog number: 389374), storage at 4 °C
8. Accutase (Corning, catalog number: 25058CI), storage at -20 °C
9. Phosphate-buffered saline (PBS) (Gibco, catalog number: 14190-094), storage at 4 °C
Solutions
1. Cell culture medium (see Recipes)
Recipes
1. Cell culture medium
Prepare the complete NBM (cNBM) and the complete NBMW/O (cNBMW/O) by adding 10 mL of B27, 20 ng/mL of basic FGF, 100 U/μL of heparin, and 1,000 U/mL of penicillin-streptomycin in 500 mL of medium. Supplement the cNBMW/O with 4.5 mg/mL of [13C]6-glucose ([13C]6-glucose NBM).
Laboratory supplies
1. Filter membrane 0.2 μm (Sartolon Polyamide)
2. Qiagen RNeasy Mini kit (Qiagen, catalog number: 74104)
Equipment
1. CO2 incubator (Panasonic, model: MCO-18AC-PE, catalog number: 13040182)
2. Hypoxia chamber (Heraeus, model: B 5060 EK/CO2, catalog number: 8403096)
3. Vacuum pump (KNF Neuberger, model: N86 KN.18, catalog number: D-79112)
Software and datasets
1. The datasets and software used here are listed in Table 1. The DIMet tool [6] and its companion tool for data formatting, TraceGroomer, are both available through two user-friendly Galaxy web platforms: Galaxy EU [15] and Workflow4Metabolomics (W4M) [16,17]. For this protocol, W4M was chosen because it includes an isotopologue correction tool [18], which is compatible with DIMet and TraceGroomer. No coding skills are required from the user.
Table 1. Software and datasets for data analysis. DIMet and TraceGroomer Galaxy versions are available through Workflow4Metabolomics (W4M) https://workflow4metabolomics.usegalaxy.fr/ and Galaxy EU (https://usegalaxy.eu/).
Type Software/dataset/resource Version Date License Access
Data https://zenodo.org/records/13741706 0.3 10/09/2024 CC BY free
Software DIMet in its web Galaxy version (source code: https://github.com/cbib/DIMet) 0.2.4 22/03/2024 MIT free
Software Tracegroomer in its web Galaxy version (https://github.com/cbib/TraceGroomer) 0.1.4 06/08/2024 MIT free
Workflow manager Galaxy Workflow4Metabolomics (W4M) https://workflow4metabolomics.usegalaxy.fr/ 23.0 06/06/2023 GPL-3.0 free
Procedure
Cells were grown as spheroids and cultured in cNBM at 37 °C in a 5% CO2 incubator. Prior to use, these cell lines were tested for mycoplasma contamination. The absence of mycoplasma was confirmed using specific PCR primers. All cell lines used for experimental procedures were cultured between passages 10 and 30.
A. Cell passage
1. Collect spheroids in a 15 mL tube.
2. Centrifuge the tube at 1,000× g for 5 min.
3. Remove the supernatant and wash with 5 mL of PBS twice.
4. Centrifuge the tube at 1,000× g for 5 min.
5. Remove the supernatant.
6. Dissociate spheroids with 1 mL of Accutase (700 U/mL) for 15 min at 37 °C.
7. Recover cells with 9 mL of [13C]6-glucose NBM.
8. Count the cells.
B. Labeled substrate administration
1. Seed 500,000 P3 cells in 6-well plates with [13C]6-glucose NBM.
2. Incubate cells for 0, 24, and 48 h in normoxia (21% O2) or in hypoxia (0.1% O2).
C. Cells extracts for metabolomics data acquisition
1. For cells in suspension (fast filtration method):
a. Drop 1 mL of cell culture on a filter membrane with 0.2 μm.
b. Rinse the filter with 2 mL of PBS.
c. Quickly remove the filter membrane from the filtration unit, put it on an aluminum foil, and freeze it in liquid nitrogen.
d. Store the filter membrane at -80 °C.
2. For cell supernatants:
a. Recover the filtered supernatant from the filtration unit.
b. Store the supernatant at -80 °C before shipment.
3. Follow the guidance of the core services. In our case, the metabolomic profiling was performed in collaboration with MetaboHub-MetaToul.
D. RNA purification for transcriptomics data acquisition
1. Seed 500,000 P3 cells in 6-well plates with cNBM.
2. Incubate cells for 0, 24, and 48 h in normoxia (21% O2) or hypoxia (0.1% O2).
3. Freeze the cells as in section C.
4. Extract the RNA from fresh frozen cells using the Qiagen RNeasy Mini kit according to the manufacturer's protocol.
5. Check the quality and quantity of RNA. We used a fragment analyzer (Agilent) with the company's Standard Sensitivity RNA Kit (DNF‐471). Transcriptomic sequencing was performed in collaboration with the Core Unit for Molecular Tumor Diagnostics (CMTD), National Center for Tumor Diseases (NCT), Dresden, Germany.
E. Shipping of the biological samples for omics data acquisition
Send all the samples in dry ice. This is valid for both samples shipped to the metabolomics core and samples shipped to the transcriptomics platform. Metabolomics data acquisition and RNA sequencing are out of the scope of this protocol; both can be consulted in our previous publication [2].
F. Collecting the metabolomics and transcriptomics datasets
1. Collect the labeled metabolomics data. Check that the data has been corrected by the occurrence of natural isotopes; most metabolomics services provide corrected abundances of isotopologues. This correction is carried out with external tools such as IsoCor [18], available in W4M.
2. Collect the transcriptomics (RNA-seq) data, acquired by the sequencing services, and perform the differential analysis to obtain the tables of differentially expressed genes (DEG) (see Note 1).
Data analysis
A. Preparing data for the analysis with DIMet in Galaxy
1. Download the provided data from Zenodo (https://zenodo.org/records/13741706), and review its detailed description. The metabolomics data has been pre-corrected using IsoCor by the metabolomics facility. In addition to the metabolomics data, differentially expressed genes (DEG) of LDHAB-KO vs. Control, at T0 and T48 time points, are provided. Save and unzip the downloaded folder on your local machine.
2. Ensure the metabolomics data includes at least three biological replicates per group (≥ 5 biological replicates is optimal). The data provided has three biological replicates.
B. Accessing DIMet tools through the Galaxy web workflow manager
1. Open Galaxy W4M https://workflow4metabolomics.usegalaxy.fr/ in your browser. If unfamiliar with Galaxy, refer to File S1 for guidance on loading files, opening tools, and reusing files. Run all subsequent steps in the same Galaxy session for easy file reuse.
2. Only a web browser is required; no coding skills are necessary.
C. Formatting the metabolomics data with TraceGroomer
1. Open the TraceGroomer tool in Galaxy (Figure 1A) to format data for compatibility with DIMet:
a. Upload the labeled metabolomics file and the samples metadata file (Figure 1A.i).
b. Drag and drop the appropriate files to the required fields as shown in Figure 1A.ii.
c. Leave fields for optional files empty and use default settings for input type (“isocor output file”) and advanced options. Run the tool.
d. Download the output files. TraceGroomer generates four files (Figure 1A.iv) [6]. These four quantification files will be used in the following steps by the DIMet modules.
Figure 1. Format and globally explore the dataset. (A) Formatting with TraceGroomer. Upload the metadata file and the metabolomics file (i), then drag and drop them to their respective fields (ii). Run (iii). The four output files are the isotopologue abundances, the isotopologue proportions, the mean enrichment, and the total metabolite abundances (iv). (B) Explore the global data structure with the principal components analysis (PCA) plot. Drag and drop the total abundances and mean enrichment (generated in A) and the metadata to the fields as shown (i). Select the conditions (ii, upper), the output format (ii, lower-left), and run (ii, lower-right). The output consists of two types of figures: the projections of principal components 1 and 2 (scatterplots) (iii, upper left; iii, right), and the component variabilities (bars) (iii, lower-left).
D. Exploring the global data structure with the principal components analysis (PCA) using DIMet
1. Open the dimet pca plot module in Galaxy, continuing the same session used in section C.
2. Drag and drop the appropriate files to the fields, as shown in Figure 1B.i. You need to provide the total metabolite abundances (or mean enrichment) file along with the samples metadata file.
3. Set the parameters as shown in Figure 1B.ii. Select the two conditions (control and LDHAB-KO) from the drop-down menu and specify the desired output format (.pdf or .svg) using the radio buttons. Run the module.
4. Once executed, visualize the results (Figure 1B.iii) and download them (Note 2).
E. Exploring metabolite-by-metabolite quantifications using DIMet
1. For each dedicated module, run them separately (Table 2 and Figure 2A) in your session. Drag and drop the input quantification file for each module as indicated in Table 2. Continue to assign the metadata file as per the previous steps.
Table 2. DIMet modules to generate figures of individual metabolites. Each one of the three modules accepts a corresponding quantification file.
Open this module in Galaxy Drag and drop this quantification file as input
dimet enrichment plot mean enrichment
dimet isotopologues plot isotopologue proportions
dimet abundance plot total metabolite abundances
Figure 2. Explore metabolite-by-metabolite quantifications using DIMet. The horizontal panels represent the steps, whereas the vertical panels represent each one of the three modules. (A) The three modules’ names must be used separately. (B) Common parameters for the three modules: set a sequential order within the Conditions box as indicated in (i), and select all the time points (ii, upper). (C) Adjust the module-specific parameters to obtain identical output as in D. (D) Output plots. For the isotopologues (center) and the total abundance bars (right), the legend is generated as a separate file.
2. Set data-specific parameters as shown in Figure 2B, which are consistent across all three modules. Next, adjust the module-specific parameters as shown in Figure 2C.
3. Run the modules. Once completed, visualize and download the results (Figure 2D).
F. Performing the DIMet differential analysis between two groups (LDHAB-KO and control samples)
1. Open the dimet differential analysis module in Galaxy.
2. Identify the files generated by section C and the metadata within your session (Figure 3A). The dimet differential analysis accepts any type of quantification file, but only one type per run.
Figure 3. Differential analysis between two groups with DIMet. (A) Active files in the session. (B) Choose the type of quantification file in the drop-down menu (upper), then drag and drop the quantification file (center) and the metadata (lower) to their fields. (C) Select “Wilcoxon’s rank sum test” as the statistical test. (D) Define the two groups to be compared: in the Conditions box, set “LDHAB-KO” and “Control” in the indicated order (upper); in the browse time point menu, select T48 (lower). (E) Select fdr_bh (which corresponds to the Benjamini-Hochberg method) as the multiple test correction method and run. The output table (F) can be visualized and downloaded.
3. Drag and drop the input files as shown in Figure 3B.
4. Set the nonparametric statistical test as shown in Figure 3C.
5. Define the two groups to be compared, as indicated in Figure 3D. In this example, LDHAB-KO vs. control at T48 is compared. You can use any other time point to do this exercise.
6. Select the method for correcting multiple tests, as shown in Figure 3E (see Note 3), and then run the module.
7. Visualize the resulting output table (Figure 3F) and download it.
G. Performing the time-course analysis with DIMet
1. Open the dimet timecourse analysis module, which allows to compare consecutive time points.
2. Drag and drop one (any) of the quantification files as shown in Figure 3B and the metadata.
3. Select the statistical test and the method for multiple test correction as in the pairwise differential analysis (see section F).
4. In the Conditions box (Figure 3D, upper), specify the conditions that will be used for the analysis (note that conditions are not compared here).
5. Run the module. DIMet will automatically compare consecutive time points (tx+1 vs. tx) within each condition.
6. Once the results are available, download the output tables. The file names indicate the comparisons performed.
H. Comparing the entire labeling profiles between conditions or time points with DIMet
1. Open the dimet bivariate analysis module to compare entire labeling profiles (see Note 4).
2. Drag and drop the isotopologue proportions file as shown in Figure 3B and the metadata.
3. Select the Spearman test in the statistical test to apply radio buttons section.
4. In the Conditions box (Figure 3D), set the desired conditions. Then, select “fdr_bh” for multiple test correction (using radio buttons), as shown in Figure 3E.
5. Run the module. DIMet will automatically perform comparisons: (i) between LDHAB-KO and control samples for each time point, and (ii) tx+1 vs. tx for each condition.
6. Download the output tables. File names will indicate the performed comparisons.
I. Integrating metabolomics and transcriptomics data with DIMet
1. Open the dimet metabologram module to integrate labeled metabolomics with transcriptomics datasets in a pairwise differential analysis. Continue in the same session to reuse the total metabolite abundances file (generated in section C) and the metadata file (Figure 4A.i).
Figure 4. Integrate metabolomics and transcriptomics data with DIMet. (A) Files ready for the dimet metabologram module: the metabolomics data files (i), the two pathways files (ii), and the two differentially expressed genes (DEG) files (iii). (B) Drag and drop the metabolomics and pathways files (B.i, B.ii); quickly visualize the DEG files with the eye button (B.iii); identify the columns for gene symbols (lower-left) and the numeric data (lower-right). (C) Define the first comparison: drag and drop the T0 DEG file (C, upper). In the drop-down menus, choose the columns seen in B.iii and the time point T0 (which is read from the metadata) (C, center). Set the Conditions box as indicated (C, lower). (D) Define the second comparison: click the insert deregulated set button to open the fields; drag and drop the T48 DEG file, choose the time point T48, and set the rest of the fields as in C. (E) Set the compartment (upper), set the statistical parameters as shown (center, lower), and run. (F) Results are downloadable; each metabologram is generated as an independent figure, as well as the global legend. The downloaded figures were arranged (F) by using external graphics software (Inkscape). Note that “transcript” and “gene” are interchangeable terms for the metabolograms.
2. From the Zenodo download (https://zenodo.org/records/13741706) in section A, locate the two files containing metabolic pathway data and the two files of the differentially expressed genes (DEGs). These four files are found in the subfolder metabologram_data/. Upload the files to your Galaxy session as illustrated in Figures 4A.ii and 4A.iii.
3. Ensure that all active files appear as depicted in Figure 4A. Drag and drop the files to their respective fields as illustrated in Figures 4B, 4C, and 4D.
4. Set the comparisons to appear in the output (Figures 4C and 4D) and configure the global parameters (Figure 4E). Run the tool.
5. Once complete, download the results (Figure 4F).
Validation of protocol
This protocol was validated in two of our previously published papers. The first explored the role of lactate dehydrogenases in the metabolic plasticity of glioblastoma [2], with the analysis presented in Figure 4C and Appendix Figure S4. The second paper demonstrated the application of the DIMet tool to real tracer metabolomics datasets, including a time-series dataset from glioblastoma control cultures [6].
General notes and troubleshooting
General notes
To learn how to create and organize your own files for data analysis (such as sample metadata, pathway lists, DEG files, etc.), refer to the Help menu within the DIMet Galaxy modules. There, you will find detailed guidance on the expected content and file organization for each input type.
1. We processed the bulk RNA-seq data using a standard pipeline [2]. Briefly, after preprocessing and mapping the fastq files, the resulting raw count matrix was subjected to differential expression analysis, to obtain sets of differentially expressed genes (DEGs).
2. Principal components analysis can also generate numerical results (i.e., tables) using the dimet pca analysis module in Galaxy. The usage is identical to dimet pca plots.
3. For more detailed information about the statistical tests and multiple testing correction methods offered, refer to the supplementary material of our previous publication [6], section S3.
4. A metabolite’s labeling profile refers to the set of all its isotopologues, ordered by the number of labeled (carbon) atoms and expressed as proportions. This profile is also known as MDV (mass isotopomer distribution vector) [4].
Supplementary information
The following supporting information can be downloaded here:
1. File S1. Supplementary information related to general Galaxy usage
Acknowledgments
This work was funded by the PLBIO 2021 “Biologie et Sciences du Cancer” [N 221284] grant of the Institut National du Cancer (INCA), France. This protocol was first described and validated in Guyon et al. [2] and Galvis et al. [6].
Competing interests
There are no conflicts of interest or competing interests.
Ethical considerations
P3 cells were derived from patient donor samples; the regional ethical committee approved the collection of biopsy tissue at Haukeland University Hospital, Bergen, Norway (REK 013.09).
References
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Article Information
Publication history
Received: Sep 13, 2024
Accepted: Nov 20, 2024
Available online: Dec 10, 2024
Published: Jan 20, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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5,169 | https://bio-protocol.org/en/bpdetail?id=5169&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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Primary Neuronal Culture and Transient Transfection
ST Shun-Cheng Tseng
PC Peng-Tzu Chen
EH Eric Hwang
Published: Vol 15, Iss 2, Jan 20, 2025
DOI: 10.21769/BioProtoc.5169 Views: 342
Reviewed by: Chiara AmbrogioShivaprasad H. SathyanarayanaSneha Ray
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Original Research Article:
The authors used this protocol in eLIFE Jun 2024
Abstract
Primary neuronal culture and transient transfection offer a pair of crucial tools for neuroscience research, providing a controlled environment to study the behavior, function, and interactions of neurons in vitro. These cultures can be used to investigate fundamental aspects of neuronal development and plasticity, as well as disease mechanisms. There are numerous methods of transient transfection, such as electroporation, calcium phosphate precipitation, or cationic lipid transfection. In this protocol, we used electroporation for neurons immediately before plating and cationic lipid transfection for neurons that have been cultured for a few days in vitro. In our experience, the transfection efficiency of electroporation can be as high as 30%, and cationic lipid transfection has an efficiency of 1%–2%. While cationic lipid transfection has much lower efficiency than electroporation, it does offer the advantage of a higher expression level. Therefore, these transfection methods are suitable for different stages of neurons and different expression requirements.
Key features
• Culture of primary neurons from the CNS.
• Electroporation for freshly isolated neurons in suspension.
• Cationic lipid transfection for adherent neurons.
Keywords: Neuronal morphology Neurite outgrowth Neuronal isolation Adherent neuron transient transfection Suspension neuron transient transfection
Graphical overview
Background
Neurons are highly specialized cells whose development and function are influenced by complex interactions of genetic and environmental factors. In vitro culture systems offer a controlled environment for studying neuroscience at the cellular level. While primary neuronal culture has its limitations (e.g., losing the complexity of the in vivo network or the stiffness of the culture surface being different from that of the nervous system), it offers a controlled environment to assess of effect of biological, chemical, or physical perturbations. To study the physiological roles of target proteins, we need to alter the expression or sequence of the target genes in neurons through transfection. Electroporation and cationic lipid transfection are two non-viral methods for introducing exogenous DNA into primary neurons. Both methods have been widely used, and their efficacy varies significantly depending on the types and developmental stages of neurons [1].
Electroporation has the advantage when the target cells are freshly isolated neurons or cells in suspension that have yet to produce cellular protrusions [1]. Electroporation can provide high transfection efficiency [2] and can be applied to a wide range of biological studies because of its versatility [3]. Electroporation is not a time-consuming procedure, allowing for efficient transfection of large numbers of cells [4], and can directly deliver exogenous DNA into cells without viral or non-viral vectors [2]. Compared to other methods of gene delivery, electroporation is generally less toxic than others [2]. It has the potential to be utilized not only for DNA transfer but also for other charged macromolecules (such as chemical molecules, antibodies, antisense oligonucleotides, RNAs, and artificial chromosomes) into neurons [5].
However, electroporation is not a perfect method for neurons at later stages of development. Adherent neurons with neurites are highly sensitive to physical stress, including the electric pulses used in electroporation [6]. In primary neuronal cultures that have been maintained in vitro for a few days, the close proximity of somata, axons, and dendrites can alter the electric field experienced by each cell compared to low-density cell suspensions. Furthermore, the presence of extracellular matrix in cultures can further influence the electroporation process [7]. To transfect neurons that have been cultured in vitro for a few days, cationic lipid transfection is a good alternative. Cationic lipid transfection achieves reliable efficiency in adherent neurons with neurites due to its ability to form unilamellar liposomes that facilitate the entry of nucleic acids into the cells. Cationic liposomes, with diameters between 100 and 500 nm, are lipid spheres with positive charges on their surface [8], which are attracted to the negative charges of both DNA and the cell membrane of neurons. This makes cationic lipid transfection less dependent on the physical properties of the cells, making it more suitable for complex cell types [9]. Unlike electroporation, cationic lipid transfection does not involve the application of electric pulses. This reduces the physical stress on neurons and improves the survival rate of the cells [2]. Additionally, cationic lipid transfection can result in higher gene expression than electroporation in our hands.
Electroporation is a powerful tool for transfection in freshly isolated neurons because of its high efficiency, while cationic lipid transfection offers a higher gene expression in adherent neurons.
Materials and reagents
Animals
1. Pregnant (E17.5) C57BL/6NCrlBltw mice (BioLASCO Taiwan Co., C57BL/6)
Reagents
1. Fetal bovine serum (FBS) (Thermo Fisher, catalog number: 10437028)
2. B27 supplement 50× (Thermo Fisher, catalog number: 17504044)
3. Lipofectamine 2000 (LFA) (Thermo Fisher, catalog number: 11668027)
4. Minimum essential medium (MEM) (Thermo Fisher, catalog number: 11090099)
5. Neurobasal medium (NB) (Thermo Fisher, catalog number: 21103049)
6. 0.25% trypsin-EDTA (Thermo Fisher, catalog number: 25200056)
7. RPMI 1640 medium (Thermo Fisher, catalog number: 21870076)
8. Mouse Neuron Nucleofector kit (Lonza, catalog number: VPG-1001)
9. 10× Hank's balanced salt solution (HBSS) (Thermo Fisher, catalog number: 14185052)
10. 1 M N-2-hydroxyethylpiperazine-N-2-ethane sulfonic acid (HEPES) (Thermo Fisher, catalog number: 15630080)
11. Penicillin–streptomycin (10,000 U/mL) (Thermo Fisher, catalog number: 15140122)
12. D-glucose (Sigma-Aldrich, catalog number: G8769)
13. Boric acid (J.T. Baker, catalog number: 4035-01)
14. Sodium tetraborate (Sigma-Aldrich, catalog number: 31457)
15. Poly-L-lysine, MW 30,000–70,000 (Sigma-Aldrich, catalog number: P2636)
16. DNase I (Sigma-Aldrich, catalog number: DN25)
17. L-glutamine (Sigma-Aldrich, catalog number: G8540)
18. Acetone (Sigma-Aldrich, catalog number: 32201)
19. Ethanol absolute (Sigma-Aldrich, catalog number: 32221)
20. Trypan blue (Thermo Fisher, catalog number: 12250061)
Solutions
1. Boric acid buffer (see Recipes)
2. Poly-L-lysine stock solution (see Recipes)
3. Calcium- and magnesium-free HBSS (CMF-HBSS) (see Recipes)
4. DNase I solution (see Recipes)
5. Digestion medium (see Recipes)
6. L-glutamine solution (see Recipes)
7. Neuronal plating medium (see Recipes)
8. Neuronal maintenance medium (see Recipes)
Recipes
1. Boric acid buffer
Reagent Final concentration Quantity or Volume
Boric acid 50 mM 1.24 g
Sodium tetraborate 12 mM 1.90 g
Adjust to pH 8.5 with 1 M NaOH, add sterile ddH2O to 400 mL, and store at 4 °C.
2. Poly-L-lysine stock solution
Reagent Final concentration Quantity or Volume
Poly-L-lysine 1 mg/mL 50 mg
Boric acid buffer 50 mL
Use the top bottle filter to sterilize and store at 4 °C.
The working solution is made by diluting the stock solution with boric acid buffer to 100 μg/mL.
3. Calcium- and magnesium-free HBSS (CMF-HBSS)
Reagent Final concentration Quantity or Volume
1 M HEPES 10 mM 5 mL
Penicillin–streptomycin (10,000 U/mL) 100 U/mL 5 mL
10× HBSS 1× 50 mL
Add sterile ddH2O to 500 mL and store at 4 °C.
4. DNase I solution
Reagent Final concentration Quantity or Volume
DNase I 10 mg/mL 10 mg
ddH2O 1 mL
Aliquot into 1.5 mL microcentrifuge tubes and store at -20 °C.
5. Digestion medium
Reagent Final concentration Quantity or Volume
1 M HEPES 10 mM 1 mL
0.25% trypsin-EDTA 100 mL
Aliquot into 15 mL centrifuge tubes and store at -20 °C.
6. L-glutamine solution
Reagent Final concentration Quantity or Volume
L-glutamine 200 mM 1.46 g
Add ddH2O to 50 mL. Warm up in a 37 °C water bath for 30 min, vortex the tube every 5 min to ensure L-glutamine is dissolved, aliquot into 1.5 mL microcentrifuge tubes, and store at -20 °C.
7. Neuronal plating medium
Reagent Final concentration Quantity or Volume
D-glucose 0.6% 200 μL
FBS 5% 750 μL
L-glutamine solution 2 mM 150 μL
Add MEM so the final volume is 15 mL.
8. Neuronal maintenance medium
Reagent Final concentration Quantity or Volume
L-glutamine solution (200 mM) 0.5 mM 125 μL
B27 supplement 50× 1× 1 mL
Add neurobasal medium so the final volume is 50 mL. B27 is light-sensitive.
Laboratory supplies
1. Untreated polystyrene 24-well plate (Corning, catalog number: 351147)
2. 12 mm round glass coverslips (Marienfeld, catalog number: 0111520)
3. 70 μm cell strainer (Corning, catalog number: 352350)
4. 0.22 μm 500 mL PES top bottle filter (Corning, catalog number: COR431118)
5. 0.22 μm PES syringe filter (Sartorius, catalog number: 16532-k)
6. 50 mL syringe without needle (TERUMO, catalog number: 2TEM-SS50LZ/12)
7. 10 cm cell culture dish (Corning, catalog number: COR-430167)
8. 9 cm Petri dish (UR Brand, catalog number: UR-PD15R-1BOX)
9. 50 mL centrifuge tube (Corning, catalog number: 430829)
10. 15 mL centrifuge tube (Corning, catalog number: 430791)
11. 1.5 mL microcentrifuge tube (SSI Bio, catalog number: 1SSI-1260-00)
Equipment
1. Dissection microscope (Nikon, model: SMZ745)
2. Dissection tools:
a. Dumont #5 forceps (Roboz Surgical Instrument, model: RS-5045)
b. Dumont #7 forceps (Roboz Surgical Instrument, model: RS-5047)
c. Iris scissors (Dimeda Surgical Instruments, model: 08.340.11)
d. Iridectomy scissors (Dimeda Surgical Instrument, model: 09.140.08)
3. Laminar flow cabinet (Tsaohsin Enterprise, model: TH-420)
4. Biosafety cabinet (Esco, model: AC2-462)
5. Tabletop centrifuge (Kendro laboratory product, model: Legend RT)
6. Hemocytometer (Marienfeld, model: AP73811-00710)
7. Amaxa NucleofectorTM system (Lonza, model: Amaxa Nucleofector II)
8. CO2 incubator (NuAire, model: NU-8500)
9. Water bath (Firstek, model: B206-T1)
Software and datasets
1. BioRender (https://biorender.com). The following figures were created using BioRender: The graphical overview is created in BioRender by Tseng, L. (2024), https://BioRender.com/w09e536.
Procedure
A. Preparation of coverslips or 10 cm cell culture dish
1. Wash 12 mm coverslips once with acetone and twice with absolute ethanol on a shaker (90 rpm) for 15 min at room temperature.
2. After the second wash with ethanol, transfer coverslips into the laminar flow cabinet and then wash with sterile ddH2O thrice.
3. Transfer coverslips into each well of a 24-well plate. Remove ddH2O using aspiration. For 24-well plates, apply 200 μL of 100 μg/mL poly-L-lysine (PLL) solution to each well and incubate overnight at room temperature for coverslip coating. For 10 cm cell culture dishes, add 7 mL of 100 μg/mL PLL solution. Avoid UV light exposure after PLL was applied.
4. After PLL coating, wash coverslips with ddH2O twice and air dry.
5. PLL-coated coverslips or dishes can be kept in the refrigerator for up to one week.
B. Neuron preparation and culture for primary hippocampal and cortical neurons
1. Euthanize pregnant E17.5 C57BL/6NCrlBltw dams with CO2.
2. Sterilize the abdominal fur by spraying with 75% ethanol.
3. Cut the fur, exposing the skin and muscle layer.
4. Isolate the uterus with embryos from the abdominal cavity using the Dumont #5 forceps.
5. Take out embryos from the uterus using Dumont #5 forceps and remove the placentas. Keep in ice-cold CMF-HBSS in a Petri dish before the brain dissection.
6. Hold the E17.5 embryos by the neck with Dumont #5 forceps and use the tip of the Dumont #7 forceps to gently cut open the skin and the skull. Carefully cut open the skull from the nose to the back of the head and squeeze out the brain by pinching the sides of the skull.
7. Place the embryonic brains in a new Petri dish filled with ice-cold CMF-HBSS.
8. Operating under a stereomicroscope, separate brain cortexes from each brain and remove the meninges using a pair of Dumont #5 forceps.
9. Dissect hippocampi from both cortical hemispheres and remove the fimbriae on the concave side using the Dumont #5 forceps and the iridectomy scissors.
10. Transfer hippocampi and cortexes using Dumont #7 forceps into two different 15 mL centrifuge tubes containing 5 mL of digestion medium and incubate at 37 °C for 30 min.
11. Replace the digestion medium with room-temperature CMF-HBSS and let it stand for 3 min. Repeat the step twice to allow the residual digestion medium to diffuse from the tissue.
12. Apply 100 μL of DNase I solution to the hippocampi-containing CMF-HBSS and 200 μL of DNase I solution to the cortex-containing CMF-HBSS.
13. Then, triturate hippocampi 10 times with a 10 mL serological pipette and 10 times with a 10 mL serological pipette tipped with a 1,000 μL tip until the tissue dissociates. Triturate cortexes 20 times with a 10 mL serological pipette and 20 times with a 10 mL serological pipette tipped with a 1,000 μL tip until the tissue dissociates. Then, filter the tissue through the 70 μm cell strainer to remove tissue debris.
14. Centrifuge dissociated neurons at 100× g for 10 min at room temperature and discard the supernatant.
15. Dissociate the cell pellet by flicking the bottom of the microcentrifuge tube a few times and resuspending in the 37 °C prewarmed and pre-equilibrated neuronal plating medium.
16. Determine cell number using a hemocytometer in the presence of 0.2% trypan blue solution.
17. Seed dissociated neurons onto poly-L-lysine-coated coverslips (2.5 × 103 cells/cm2 for low-density cultures and 3 × 104 cells/cm2 for regular-density cultures) in 500 μL of prewarmed and pre-equilibrated neuronal plating medium.
18. After 4 hours, replace the neuronal plating medium with the prewarmed and pre-equilibrated 50% freshly made neuronal plating medium mixed with 50% conditioned medium (for low-density cultures) or neuronal maintenance medium (for regular-density cultures). To prepare the conditioned medium, seed 2.2 × 106 cortical neurons in a poly-L-lysine-coated 10 cm cell culture dish and culture for 7 days. The cell culture medium is the conditioned medium; it can be stored at -20 °C for several months.
C. Electroporation
1. Centrifuge 1 × 106 dissociated neurons at 80× g for 10 min at room temperature and remove the supernatant.
2. Carefully resuspend the neuron pellet in 100 μL of room-temperature mouse neuron nucleofector solution plus 3 μg of DNA. Then, transfer the mixture into a sterile cuvette.
3. Perform electroporation with the Nucleofector Program O-005 for mouse neuron nucleofector.
4. Add 330 μL of prewarmed and pre-equilibrated RPMI 1640 medium to a cuvette immediately after electroporation and transfer gently (without pipetting up and down) into a 1.5 mL tube for recovery (incubating for 10 min in a 37 °C CO2 incubator).
5. After the recovery step, seed neurons into each well containing the prewarmed and pre-equilibrated neuronal plating medium using the aforementioned density.
6. After 4 h, replace the neuronal plating medium with the fresh neuronal maintenance medium.
7. The protein expressed from the plasmid can usually be detected in 24 h.
D. Cationic lipid transfection
1. Transfect neurons in each well at least 48 h after seeding using Lipofectamine 2000 (LFA). In our experience, transfecting neurons 48 h after seeding causes severe cell death.
2. For one reaction, dilute 1 μg of plasmid DNA and 1 μL of LFA with 50 μL each of neurobasal medium (NB) to make the DNA/NB complex solution and the LFA/NB complex solution. This plasmid DNA-to-LFA ratio must be empirically determined for optimal transfection efficiency.
3. After 5 min, pipette the DNA/NB complex solution into the LFA/NB solution. Mix the mixture by flicking the bottom of the tube and leave to stand at room temperature for 20 min before adding it to each neuron-containing well.
4. Forty-five minutes after transfection, replace the medium with prewarmed and pre-equilibrated 50% freshly made neuronal plating medium mixed with 50% conditioned medium (for low-density cultures) or neuronal maintenance medium (for regular-density cultures).
5. The protein expressed from the plasmid can usually be detected in 24 h.
Data analysis
Neurons can be fixed at specific days in vitro (DIV) for subsequent image analyses. For transfection efficiency quantification, neurons were transiently transfected with a plasmid expressing the cytosolic EGFP and immunofluorescence stained with antibodies against neuron-specific β-III-tubulin and the transfection indicator EGFP (Figure 1). At least 2.4 × 105 neurons (or the number of neurons on four coverslips) were counted to calculate the transfection efficiency. When the mean fluorescence intensity of the EGFP signal in the soma of a particular neuron was higher than three times the standard deviation of the mean fluorescence intensity of the background, that neuron was considered transfected. For morphological analyses, we typically use the SNT plugin for ImageJ or Fiji [10].
Figure 1. Morphology and expression of EGFP in primary cortical neurons. (A) Representative image of 3 days in vitro (DIV) primary cortical neurons immunofluorescence stained with the neuron-specific β-III-tubulin antibody. (B) Representative image of 3 DIV primary cortical neurons immunofluorescence stained with β-III-tubulin (red in the merged image) and EGFP (green) antibodies. Neurons were transfected with a cytosolic EGFP-expressing plasmid on 2 DIV using Lipofectamine 2000. Scale bars represent 50 μm.
Validation of protocol
This protocol or parts of it have been used and validated in the following research article(s):
• Chen et al. [11]. The non-mitotic role of HMMR in regulating the localization of TPX2 and the dynamics of microtubules in neurons. eLife 13. https://doi.org/10.7554/eLife.94547
General notes and troubleshooting
General notes
1. For quality control, the number of harvested cells should be consistent, with approximately 1 × 107 cortical neurons from three embryonic mouse brains.
2. For optimal trypsin digestion, the number of cortices per tube should be between 3 and 4 to prevent incomplete trypsin digestion.
3. Generally, healthy neurons should adhere to the plate within 2 h after seeding, and structures similar to lamellipodia and filopodia should appear approximately 4 h later.
4. After electroporation, the survival rate of neurons is approximately 50%. As a result, one needs to increase the seeding density to compensate for this.
5. The DNase I solution should be prepared on the same day of the experiment and stored in a 4 °C refrigerator before the experiment.
6. The entire dissection procedure should be performed in ice-cold CMF-HBSS.
7. Neuronal plating medium, neuronal maintenance medium, and RPMI 1640 medium should be placed in a CO2 incubator for at least 15 min for pH pre-equilibrium.
8. Adding the conditioned medium promotes neurite outgrowth and axonal formation. Those who are studying the morphogenetic process of neurons should be aware of this.
9. To reduce the possibility of contamination between tissues, it is recommended that separate surgical instruments be used for different dissection steps.
Troubleshooting
Problem 1: During dissection, the anatomical structure of the hippocampus is unclear, making it difficult to distinguish and separate it from the surrounding tissues.
Possible cause: Insufficient gestation period.
Solution: Confirm the calculation method of the mouse gestation period with the supplier and record data such as the number and size of embryos.
Problem 2: Dissociated cells do not completely precipitate after the centrifugation step.
Possible cause: Tissue dissociation is incomplete.
Solution: Invert the centrifuge tube to facilitate the dissociation of cells and centrifuge again.
Problem 3: Neurons are not completely dissociated after seeding.
Possible cause: Cells were not completely dissociated before replacing the digestion medium with neuronal plating medium after centrifugation.
Solution: Try tapping the centrifuge tube to disperse the cell pellet after removing the digestion medium.
Problem 4: The somata are clustered, and the neurites are bundled together.
Possible cause: Problem with PLL coating or the glass coverslip.
Solution: The quality of the glass is crucial; use borosilicate glass for the coverslip. Wash the coverslips thoroughly and do not store washed coverslips for too long before use. Make sure to use the PLL with proper molecular weight (30,000–70,000) and use the PLL-coated coverslip within one week.
Acknowledgments
This work was supported by a grant from National Science and Technology Council (NSTC 111-2320-B-A49-015-MY3) as well as Center for Intelligent Drug Systems and Smart Bio-devices (IDS2B) from the Featured Areas Research Center Program within the framework of the Higher Education Sprout Project by the Ministry of Education in Taiwan. This protocol was adapted and modified from Chen et al. [11].
Competing interests
The authors declare no competing interests.
Ethical considerations
All animal experimental procedures were approved by the Institutional Animal Care and Use Committee (IACUC) and in accordance with the Guide for the Care and Use of Laboratory Animals of National Yang Ming Chiao Tung University.
References
Karra, D. and Dahm, R. (2010). Transfection Techniques for Neuronal Cells: Table 1. J Neurosci. 30(18): 6171–6177.
Washbourne, P. and McAllister, A. (2002). Techniques for gene transfer into neurons. Curr Opin Neurobiol. 12(5): 566–573.
Shigekawa, K. and Dower, W. J. (1988). Electroporation of eukaryotes and prokaryotes: a general approach to the introduction of macromolecules into cells. Biotechniques. 6(8): 742–751.
Geisler, C. and Wirth, T. (2006). Electroporation: a simple, versatile tool for gene transfer. In: Methods in Molecular Biology. 340: 3–15.
Inoue, T. and Krumlauf, R. (2001). An impulse to the brain—using in vivo electroporation. Nat Neurosci. 4: 1156–1158.
Marwick, K. F. M. and Hardingham, G. E. (2017). Transfection in Primary Cultured Neuronal Cells. In: Methods in Molecular Biology. 1677: 137–144.
Kranjc, M. and Miklavčič, D. (2017). Electric Field Distribution and Electroporation Threshold. In: Miklavčič, D. (Ed.). Handbook of Electroporation: 1043–1058.
Pedroso de Lima, M. C., Simões, S., Pires, P., Faneca, H. and Düzgüneş, N. (2001). Cationic lipid–DNA complexes in gene delivery: from biophysics to biological applications. Adv Drug Delivery Rev. 47: 277–294.
Yu, X., Xiao, H., Muqier, M., Han, S. and Baigude, H. (2022). Effect of the array of amines on the transfection efficiency of cationic peptidomimetic lipid molecules into neural cells. RSC Adv. 12(33): 21567–21573.
Arshadi, C., Günther, U., Eddison, M., Harrington, K. I. S. and Ferreira, T. A. (2021). SNT: a unifying toolbox for quantification of neuronal anatomy. Nat Methods. 18(4): 374–377.
Chen, Y. J., Tseng, S. C., Chen, P. T. and Hwang, E. (2024). The non-mitotic role of HMMR in regulating the localization of TPX2 and the dynamics of microtubules in neurons. eLife. 13: RP94547.
Article Information
Publication history
Received: Sep 21, 2024
Accepted: Nov 27, 2024
Available online: Dec 19, 2024
Published: Jan 20, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
How to cite
Category
Neuroscience > Development > Neuron
Cell Biology > Cell isolation and culture > Monolayer culture
Molecular Biology > DNA > Transfection
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Identification of Mycobacterium tuberculosis and its Drug Resistance by Targeted Nanopore Sequencing Technology
CT Chen Tang
FX Feng Xu
XZ Xiaoqun Zheng
GX Guangxin Xiang
Published: Vol 15, Iss 3, Feb 5, 2025
DOI: 10.21769/BioProtoc.5170 Views: 48
Reviewed by: Alka MehraSoumya MoonjelySuresh Panthee
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Original Research Article:
The authors used this protocol in Frontiers in Microbiology May 2024
Abstract
Tuberculosis (TB) remains the leading cause of human mortality in infectious diseases. Drug-resistant TB, particularly multidrug-resistant TB and extensively drug-resistant TB, poses a pressing clinical and public health challenge. The main causative agents of TB are known as Mycobacterium tuberculosis (MTB), which exhibits a highly complex drug resistance profile. Traditional culture-based phenotypic drug susceptibility testing is time-consuming, and PCR-based assays are restricted to detecting known mutational hotspots. In this study, we present a protocol leveraging high-throughput nanopore sequencing technology in conjunction with multiplex PCR, termed targeted nanopore sequencing, for the identification of MTB and analysis of its drug resistance. Our method for MTB drug resistance assessment offers the benefits of being culture-free, efficient, high-throughput, and highly accurate, which could significantly aid in clinical patient management and the control of TB infections.
Key features
• Targeted nanopore sequencing detects 18 genes simultaneously linked to antibiotic resistance in MTB.
• The method provides broad drug resistance profiles for 14 first- and second-line anti-TB drugs without bacterial culture.
• The expedited turnaround time of the process is approximately 7.5 h with a detection limit of 102 bacteria/mL.
Keywords: Mycobacterium tuberculosis Drug resistance Nanopore sequencing Targeted sequencing Multiplex PCR
Graphical overview
Targeted nanopore sequencing for the drug resistance assay of Mycobacterium tuberculosis. Initially, DNA is extracted, and the targeted regions within genes associated with drug resistance are amplified via multiplex PCR. Subsequently, the amplicons undergo barcoding and adapter ligation. The library preparation is then sequenced on a nanopore sequencer, which yields long-read sequence data. Ultimately, drug resistance is identified through bioinformatics analysis. The entire targeted nanopore sequencing process has an expedited turnaround time of approximately 7.5 h.
Background
Tuberculosis (TB) affirms its position as the preeminent cause of human death from infectious diseases, with an estimated 1.7 million deaths occurring globally each year, as reported by Daley [1]. The latest figures from the World Health Organization’s Global Tuberculosis Report 2023 reveal that 7.5 million new cases were diagnosed worldwide in 2022, surpassing pre-COVID levels. TB pathogen refers to a group of bacteria within the Mycobacterium tuberculosis complex (MTBC), including Mycobacterium tuberculosis (MTB), noted for its intricate drug resistance spectrum [2]. MTB, being the principal causative agent of TB, gains antibiotic resistance through gene mutations, significantly influencing clinical treatment approaches [3–5].
Surveillance of highly virulent and multidrug-resistant (MDR) MTB strains is imperative for securing precise diagnoses and potent treatment regimens [6]. Conventional culture-based phenotypic drug susceptibility testing (pDST) is the standard method for assessing MTB, although being time-consuming, typically requiring a span of days to weeks [3]. The prevalent molecular detection methods, including the real-time PCR-based Xpert MTB/RIF assay endorsed by the WHO, while being efficient, are limited to detecting known mutational hotspots and thus provide incomplete data on drug resistance [7]. Consequently, infections with MDR and extensively drug-resistant (XDR) MTB strains highlight an urgent need for advanced diagnostic tools to guide TB treatment options [8].
High-throughput sequencing technology has become a staple in clinical diagnostics. Nanopore sequencing, a cutting-edge method exemplified by the equipment from Oxford Nanopore Technologies (ONT), offers the benefits of long-read lengths, real-time data acquisition, and rapid sequencing [9]. While its accuracy has been a subject of improvement, significant strides in scientific and technological advancements have enhanced its performance in recent years [10,11]. Nanopore sequencing facilitates direct analysis of DNA or RNA sequences without the need for culturing MTB, enabling swift identification of MTB and detection of TB drug resistance. This approach might not only reduce the overall cost of MTB detection but also expedite the diagnostic process [12]. Therefore, the technology holds considerable promise for clinical applications and possesses a promising market outlook, making it an attractive option for clinical settings [13].
This study introduces a method that integrates multiplex PCR for targeted enrichment with nanopore sequencing, termed targeted nanopore sequencing. Furthermore, the bioinformatic tool TBProfiler is employed for drug resistance analysis, with its database encompassing 1,195 candidate mutations, which were obtained globally from more than 17,000 clinical isolates and assessed with whole-genome sequencing (WGS) and pDST [14].
Materials and reagents
Reagents
1. 10× TBE buffer (Solarbio, catalog number: T1051)
2. 1 M Tris-HCl (pH 8.0) (Solarbio, catalog number: T1150)
3. 200 bp DNA ladder (Dye Plus) (Takara, catalog number: 3423A)
4. Agarose (Mei5 Biotechnology, Co., Ltd, catalog number: MF103)
5. Agencourt AMPure XP beads (Beckman Coulter, catalog number: A63881)
6. Chelex 100 sodium (Solarbio, catalog number: C8230)
7. DNase/RNase-free water (Solarbio, catalog number: R1600)
8. dNTP mixture (Takara, catalog number: 4030)
9. EDTA (Sigma, catalog number: 798681)
10. Ligation Sequencing kit (Q20+) (Oxford Nanopore Technologies, catalog number: SQK-LSK114)
11. M5 6× DNA electrophoresis loading buffer (Mei5 Biotechnology, Co., Ltd, catalog number: MF144-01)
12. M5 Hipure Next III Gelred (Mei5 Biotechnology, Co., Ltd, catalog number: MF380-01)
13. NaOH (Sigma, catalog number: 221465)
14. Native Barcoding kit 24 (Q20+) (Oxford Nanopore Technologies, catalog number: SQK-NBD114.24)
15. NEB Blunt/TA ligase master mix (New England Biolabs, catalog number: M0367L)
16. NEB Q5 Hot Start High-Fidelity DNA Polymerase (New England Biolabs, catalog number: M0493L)
17. NEBNext Ultra II end repair/dA-tailing module (New England Biolabs, catalog number: E7546S)
18. NEBNext quick ligation module (New England Biolabs, catalog number: E6056S)
19. Nonidet P40 substitute (NP-40) (Solarbio, catalog number: N8030)
20. PBS (Sigma, catalog number: P3813)
21. Qubit dsDNA HS Assay kit (Thermo Fisher Scientific, catalog number: Q32854)
22. Triton X-100 (Solarbio, catalog number: T8200)
Laboratory supplies
1. 0.2 mL PCR tubes (ideally in strips of 8) (Thermo Fisher Scientific, catalog number: AB0490)
2. 1.5 mL tubes (Thermo Fisher Scientific, catalog number: S348903)
3. Pipette tips (Thermo Fisher Scientific, catalog numbers: 02-707-426, 02-707-403, 02-707-438)
4. PromethION flow cell R10.4.1 (Oxford Nanopore Technologies, catalog number: FLO-PRO114M)
5. Qubit assay tubes (Thermo Fisher Scientific, catalog number: Q32856)
Equipment
1. Biological safety cabinet (NuAire Lab Equipment, catalog number: NU-425-300)
2. Electrophoresis unit of power supply (Thermo Fisher Scientific, catalog number: S65533Q)
3. Magnetic stand for 1.5 mL tubes (Beckman Coulter, catalog number: A29182)
4. Microcentrifuge (Eppendorf, catalog number: EP5401000137)
5. Milli-Q ultrapure water system (Millipore Synergy, catalog number: F1CA45528 A)
6. NanoDrop spectrophotometer (Thermo Fisher Scientific, catalog number: ND-1000)
7. Qubit 4 fluorometer (Thermo Fisher Scientific, catalog number: Q33238)
8. Sequencing device (Oxford Nanopore Technologies, model: PromethION)
9. S1000 thermal cycler (Bio-Rad, catalog number: 1852196)
10. Vortex mixer (Thermo Fisher Scientific, catalog number: S96461A)
Software and datasets
1. Albacore (https://github.com/Albacore/albacore)
2. Bowtie2 (https://github.com/BenLangmead/bowtie2)
3. Delly (https://github.com/dellytools/delly)
4. MinKNOW (https://github.com/nanoporetech/minknow_api)
5. NanoFilt (https://github.com/wdecoster/nanofilt)
6. SAMtools (https://github.com/samtools/samtools)
7. TBProfiler (https://github.com/jodyphelan/TBProfiler)
8. Trimmomatic (https://github.com/usadellab/Trimmomatic)
Procedure
A. DNA extraction
1. Pre-treat clinical samples such as sputum and bronchoalveolar lavage fluid with 4% NaOH and PBS solution.
2. Collect 1 mL of the sample liquid into a 1.5 mL tube and centrifuge it at 11,000× g for 5 min; then, remove 950 μL of the supernatant.
3. Add 50 μL of nucleic acid extraction reagent (1 mM EDTA, 10 mM Tris-HCl, 1% NP-40, 1% Triton X-100, and 50% Chelex 100) to the tube and vortex the mixture for 1 min. The role of Chelex 100 is to capture divalent cations from biological samples and inhibit the activity of metallonucleases, even in highly concentrated salt solutions.
4. Place the mixture tube in a metal bath and boil it at 100 °C for 10 min.
5. After boiling, vortex the tube for 1 min and then centrifuge it at 11,000× g for 5 min.
6. Take the supernatant, enriched in DNA, as the template for multiplex PCR.
B. Multiplex PCR
1. Prepare a primer mixture containing primers designed for 18 drug resistance–associated genes, namely gyrB, gyrA, rpoB, mmpR5, rpsL, rplC, atpE, rrs, rrl, fabG1, inhA, rpsA, tlyA, katG, pncA, eis, embB, and ubiA, with their primer sequences and concentrations described in our previous publication [15].
2. Combine 1 μL of the multi-primer mixture, 0.25 μL of Q5 Hot-Start High-Fidelity DNA Polymerase, 5 μL of High GC enhancer, 5 μL of reaction buffer, 2 μL of dNTPs (2.5 mM), 4 μL of DNase/RNase-free water, and 10 μL of DNA template. Mix by flicking and spin down.
3. Perform the PCR with an initial denaturation step at 98 °C for 2 min, followed by 40 cycles of denaturation at 95 °C for 25 s, annealing at 60 °C for 30 s, and extension at 72 °C for 3 min, concluding with a final extension step at 72 °C for 4 min.
4. The amplicons can be visualized by agarose gel electrophoresis and quantified using a Qubit assay.
C. Library preparation
1. Purification of PCR amplicons
a. Take the AMPure XP beads out of the refrigerator at 4 °C, vortex to mix them evenly, and allow them to equilibrate at room temperature (RT) for at least 30 min before use.
b. Adjust the volume of 24 PCR amplicon samples to the lowest concentration observed, then add equal volumes to 24 × 1.5 mL tubes, supplementing with nuclease-free water to a final volume of 24 μL per tube.
c. To each of the 24 tubes, add 36 μL (1.5×) of the AMPure XP beads for DNA purification purposes, mix by pipetting 10 times, and incubate at RT for 15 min.
d. Place the tubes on a magnetic stand for 2–3 min, then carefully remove and discard the supernatant, ensuring the beads are not disturbed.
e. Gently add 200 μL of freshly prepared 80% ethanol to each tube, incubate at RT for 30–60 s, then remove and discard the ethanol. Air dry the beads for 2–3 min and repeat the process twice.
f. Remove the tubes from the magnetic stand, resuspend beads in 24 μL of nuclease-free water, and mix gently by flicking or pipetting up and down five times. Incubate for 5 min at RT.
g. Place the tubes back on the magnetic stand and carefully transfer 22.4 μL of the supernatant containing the eluted DNA to new tubes after the solution clears.
h. Perform quantification using a Qubit assay.
2. DNA end repair
a. Add 1.4 μL of Ultra II end-prep reaction buffer and 1.2 μL of Ultra II end-prep enzyme mix (from NEBNext end repair module) to the tubes containing supernatant with eluted DNA to achieve repaired DNA with 5' phosphorylated, 3' dA-tailed ends.
b. Gently mix the solution by pipetting up and down five times and incubate at 20 °C for 50 min, followed by a 20-min incubation at 65 °C in a thermocycler.
c. Follow the same bead addition and incubation steps as in steps C1c, d, and e.
d. Resuspend the beads in 16 μL of nuclease-free water, mix gently, and incubate for 5 min at RT.
e. Place the tubes back on the magnetic stand and carefully transfer 14.5 μL of the supernatant with eluted DNA to new tubes after the solution clears.
f. Perform quantification using a Qubit assay.
3. Ligation of barcodes
a. Add 1.5 μL of barcodes (Native Barcoding kit 24) and 15 μL of Blunt/TA ligase (NEB Blunt/TA ligase master mix) to the tubes with supernatant containing eluted DNA, providing 24 unique barcodes to enable multiplexing of amplicons.
b. Gently mix the solution and incubate at 20 °C for 60 min in a thermocycler, mixing every 20 min.
c. Follow the same bead addition and incubation steps as in steps C1c, d, and e.
d. Resuspend the beads in 10 μL of nuclease-free water, mix gently, and incubate for 5 min at RT.
e. Place the tubes back on the magnetic stand and carefully transfer 10 μL of the supernatant with eluted DNA to new tubes after the solution clears.
f. Perform quantification using a Qubit assay.
4. Ligation of sequencing adapters
a. Combine 24 PCR products into a new tube to create a library pool with an equal amount of DNA.
b. To the 25 μL library pool, add 10 μL of ligation buffer, 5 μL of NEBNext Quick T4 DNA ligase, and 5 μL of native adapter from the Oxford Nanopore Ligation Sequencing kit. Mix gently and briefly spin down the reaction.
c. Incubate the library pool for 60 min at 20 °C in a thermocycler.
d. Follow the same bead addition and incubation steps as in steps C1c, d, and e.
e. Add 200 μL of short fragment buffer to the library pool, incubate for 1 min, and carefully discard the supernatant. Repeat the process twice.
f. Resuspend the beads in 22 μL of elution buffer provided in the Ligation Sequencing kit and mix gently.
g. Spin down the pool and incubate for 10 min at RT.
h. Place the pool back onto the magnetic stand for 1 min and then transfer the eluate with the DNA library to a new tube, discarding the beads.
i. Perform quantification using a Qubit assay; the concentration is approximately 10 ng/μL.
D. Nanopore sequencing
1. Combine 30 μL of flow cell tether (Ligation Sequencing kit) with 1,170 μL of flow cell flush to prepare the priming mix, which is used for initiating flow cells with enough active pores for sequencing.
2. Apply 600 μL of the priming mix to the PromethION flow cell (R10.4.1) with R10 nanopores for high-consensus accuracy sequencing, wait for 5 min, and then add the rest of the mix.
3. Mix 21 μL of the library, 34 μL of loading beads, and 50 μL of sequencing buffer by gently inverting the tube 10 times.
4. Load 105 μL of the prepared library into the PromethION flow cell (R10.4.1) and initiate DNA sequencing for 1 h as per the manufacturer's protocol.
E. Bioinformatics analysis
1. Primary data acquisition is performed using MinKNOW software, which operates nanopore sequencing devices with a feature-rich user interface and saves data in FAST5 file format.
2. Albacore conducts base recognition to convert data from FAST5 to raw sequence data in FASTQ file format.
3. Reads with a Phred quality value ≤ 9 are filtered out by NanoFilt.
4. The bioinformatics pipeline TBProfiler detects antibiotic resistance mutations and can be accessed from its GitHub repository (https://github.com/jodyphelan/TBProfiler), installed via Bioconda, and utilized for resistance analysis. In Phelan et al. [14], a schematic was provided, highlighting the main steps in the TBProfiler pipeline, as well as an example of the TBProfiler report, illustrating the final output.
a. Compile sequencing read datasets from public MTB genomes with known resistance profiles for both first- and second-line antibiotics.
b. Update the TBProfiler mutation library as needed.
c. In standard operation, reads are trimmed with Trimmomatic (parameters: LEADING:3, TRAILING:3, SLIDINGWINDOW:4:20, MINLEN:36) and aligned to the H37Rv reference genome (GenBank accession no. NC_000962.3) using Bowtie2 with default parameters.
d. Call variants with BCFtools mpileup (parameters: -ABq0, -Q0, -a DP, AD) and BCFtools call (parameters: -mg 10), annotate with BCFtools csq (parameters: -p m), and parallelize the process with GNU parallel. Deletion calling is performed using Delly.
e. TBProfiler generates reports in JSON, TXT, and PDF formats, organizes data for visualization on a phylogenetic tree with iTOL, and allows for the generation and upload of config files to iTOL for visualizing drug resistance types, lineage, and individual drug resistance predictions.
Validation of protocol
Targeted nanopore sequencing, an advanced method that integrates nanopore sequencing technology with multiplex PCR, is capable of sequencing 18 genes simultaneously linked to antibiotic resistance in MTB in approximately 7.5 h. This method not only differentiates MTB from other bacterial species but also achieves a detection limit of 102 bacteria/mL by analyzing the datasets of all 18 targeted genes from 102 bacteria/mL to 107 bacteria/mL. Additionally, it furnishes detailed drug resistance profiles for 14 first- and second-line anti-TB drugs, facilitating informed drug prescription (see Graphical overview). While PromethION was utilized in this context, alternative devices like the portable MinION could be employed. Comprehensive data analysis and discussion are available in the publication by Tang et al. [15], accessible through the provided DOI: https://doi.org/10.3389/fmicb.2024.1331656.
General notes and troubleshooting
1. In this protocol, a single-tube multiplex PCR is developed for simultaneous amplification of 18 genes, with the set of PCR primers screened and validated.
2. The read depth of each gene cannot be consistent across the 18 targeted genes, and by optimizing the sequence and the concentration of each primer pair in the primer mixture, the output data can achieve better performance.
Acknowledgments
The authors thank the Scientific Research Center of Wenzhou Medical University for consultation and instrument availability, the Department of Clinical Laboratory from Wenzhou Central Hospital for providing clinical samples, and the original research paper [15] in which this protocol was described and validated. This research was supported by the Zhejiang Provincial Natural Science Foundation of China (Grant No. LTGG24H200001) and the Research Initiation Fund of Wenzhou Medical University (Grant No. GTJ21023).
Competing interests
The authors declare that they have no conflicts of interest with the contents of this article.
Ethical considerations
The study design was approved by the Research Ethics Board of Wenzhou Central Hospital, and all analyses were performed following the Declaration of Helsinki (L2023-02-033).
References
Daley, C. L. (2019). The Global Fight Against Tuberculosis. Thorac Surg Clin. 29(1): 19–25.
Gagneux, S. (2018). Ecology and evolution of Mycobacterium tuberculosis. Nat Rev Microbiol. 16(4): 202–213.
Koch, A., Cox, H. and Mizrahi, V. (2018). Drug-resistant tuberculosis: challenges and opportunities for diagnosis and treatment. Curr Opin Pharmacol. 42: 7–15.
Singh, V. and Chibale, K. (2021). Strategies to Combat Multi-Drug Resistance in Tuberculosis. Acc Chem Res. 54(10): 2361–2376.
Salari, N., Kanjoori, A. H., Hosseinian-Far, A., Hasheminezhad, R., Mansouri, K. and Mohammadi, M. (2023). Global prevalence of drug-resistant tuberculosis: a systematic review and meta-analysis. Infect Dis Poverty. 12(1): 57.
Consortium, C., Allix-Beguec, C., Arandjelovic, I., Bi, L., Beckert, P., Bonnet, M., Bradley, P., Cabibbe, A. M., Cancino-Munoz, I., Caulfield, M. J., et al. (2018). Prediction of susceptibility to first-line tuberculosis drugs by DNA sequencing. N Engl J Med. 379(15): 1403–1415.
Weinrick, B. (2020). Genotyping of Mycobacterium tuberculosis Rifampin Resistance-Associated Mutations by Use of Data from Xpert MTB/RIF Ultra Enables Large-Scale Tuberculosis Molecular Epidemiology Studies. J Clin Microbiol. 58(1): e01504–19.
Wu, X., Liang, R., Xiao, Y., Liu, H., Zhang, Y., Jiang, Y., Liu, M., Tang, J., Wang, W., Li, W., et al. (2023). Application of targeted next generation sequencing technology in the diagnosis of Mycobacterium Tuberculosis and first line drugs resistance directly from cell-free DNA of bronchoalveolar lavage fluid. J Infect. 86(4): 399–401.
Shendure, J., Balasubramanian, S., Church, G. M., Gilbert, W., Rogers, J., Schloss, J. A. and Waterston, R. H. (2017). DNA sequencing at 40: past, present and future. Nature. 550(7676): 345–353.
Gómez-González, P. J., Campino, S., Phelan, J. E. and Clark, T. G. (2022). Portable sequencing of Mycobacterium tuberculosis for clinical and epidemiological applications. Briefings Bioinf. 23(5): e1093/bib/bbac256.
Sereika, M., Kirkegaard, R. H., Karst, S. M., Michaelsen, T. Y., Sørensen, E. A., Wollenberg, R. D. and Albertsen, M. (2022). Oxford Nanopore R10.4 long-read sequencing enables the generation of near-finished bacterial genomes from pure cultures and metagenomes without short-read or reference polishing. Nat Methods. 19(7): 823–826.
Dippenaar, A., Goossens, S. N., Grobbelaar, M., Oostvogels, S., Cuypers, B., Laukens, K., Meehan, C. J., Warren, R. M. and van Rie, A. (2022). Nanopore Sequencing for Mycobacterium tuberculosis: a Critical Review of the Literature, New Developments, and Future Opportunities. J Clin Microbiol. 60(1): e00646–21.
Hu, T., Chitnis, N., Monos, D. and Dinh, A. (2021). Next-generation sequencing technologies: An overview. Hum Immunol. 82(11): 801–811.
Phelan, J. E., O’Sullivan, D. M., Machado, D., Ramos, J., Oppong, Y. E. A., Campino, S., O’Grady, J., McNerney, R., Hibberd, M. L., Viveiros, M., et al. (2019). Integrating informatics tools and portable sequencing technology for rapid detection of resistance to anti-tuberculous drugs. Genome Med. 11(1): 41.
Tang, C., Wu, L., Li, M., Dai, J., Shi, Y., Wang, Q., Xu, F., Zheng, L., Xiao, X., Cai, J., et al. (2024). High-throughput nanopore targeted sequencing for efficient drug resistance assay of Mycobacterium tuberculosis. Front Microbiol. 15: e1331656.
Article Information
Publication history
Received: Jul 4, 2024
Accepted: Nov 28, 2024
Available online: Dec 15, 2024
Published: Feb 5, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Category
Microbiology > Microbial genetics > DNA
Molecular Biology > DNA > DNA sequencing
Microbiology > Antimicrobial assay > Antibacterial assay
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High-resolution Cryo-EM Structure Determination of a-Synuclein—A Prototypical Amyloid Fibril
JS Juan C. Sanchez
JP Joshua A. Pierson
CB Collin G. Borcik
CR Chad M. Rienstra
EW Elizabeth R. Wright
Published: Vol 15, Iss 3, Feb 5, 2025
DOI: 10.21769/BioProtoc.5171 Views: 114
Reviewed by: Neha NandwaniVamseedhar RayaproluSuresh Kumar
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Cited by
Original Research Article:
The authors used this protocol in Nature Communications Mar 2024
Abstract
The physiological role of a-synuclein (a-syn), an intrinsically disordered presynaptic neuronal protein, is believed to impact the release of neurotransmitters through interactions with the SNARE complex. However, under certain cellular conditions that are not well understood, a-syn will self-assemble into β-sheet-rich fibrils that accumulate and form insoluble neuronal inclusions. Studies of patient-derived brain tissues have concluded that these inclusions are associated with Parkinson’s disease, the second most common neurodegenerative disorder, and other synuclein-related diseases called synucleinopathies. In addition, repetitions of specific mutations to the SNCA gene, the gene that encodes a-syn, result in an increased disposition for synucleinopathies. The latest advances in cryo-EM structure determination and real-space helical reconstruction methods have resulted in over 60 in vitro structures of a-syn fibrils solved to date, with a handful of these reaching a resolution below 2.5 Å. Here, we provide a protocol for a-syn protein expression, purification, and fibrilization. We detail how sample quality is assessed by negative stain transmission electron microscopy (NS-TEM) analysis and followed by sample vitrification using the Vitrobot Mark IV vitrification robot. We provide a detailed step-by-step protocol for high-resolution cryo-EM structure determination of a-syn fibrils using RELION and a series of specialized helical reconstruction tools that can be run within RELION. Finally, we detail how ChimeraX, Coot, and Phenix are used to build and refine a molecular model into the high-resolution cryo-EM map. This workflow resulted in a 2.04 Å structure of a-syn fibrils with excellent resolution of residues 36–97 and an additional island of density for residues 15–22 that had not been previously reported. This workflow should serve as a starting point for individuals new to the neurodegeneration and structural biology fields. Together, this procedure lays the foundation for advanced structural studies of a-syn and other amyloid fibrils.
Key features
• In vitro fibril amplification method yielding twisting fibrils that span several micrometers in length and are suitable for cryo-EM structure determination.
• High-throughput cryo-EM data collection of neurodegenerative fibrils, such as alpha-synuclein.
• Use of RELION implementations of helical reconstruction algorithms to generate high-resolution 3D structures of a-synuclein fibrils.
• Brief demonstration of the use of ChimeraX, Coot, and Phenix for molecular model building and refinement.
Keywords: Cryo-EM Helical reconstruction Alpha-synuclein Amyloid proteins Neurodegeneration Vitrification
Graphical overview
Graphical overview of a-synuclein fibrilization and cryo-EM structure determination. a-syn protein expression and purification is followed by a fibrilization protocol yielding twisting filaments that span several micrometers in length and are validated by negative stain transmission electron microscopy (NS-TEM). The sample is then vitrified, followed by cryo-EM data collection. Real-space helical reconstruction is performed in RELION to generate an electron potential map that is used for model building.
Background
Amyloid formation within neurons has been well documented to cause neurodegeneration in patients leading to a variety of diseases including Alzheimer’s (AH), Parkinson’s disease (PD), Lewy Body disease (LB), and multiple system atrophy (MSA) [1–3]. The formation of amyloids is due to protein aggregation resulting in helical, filamentous assemblies with cross β-sheet quaternary structure (Figure 1) [4]. Amyloid filaments interact with different cellular components such as membranes, cytoskeletal factors, and other filaments to form inclusion bodies that disrupt cellular processes and ultimately lead to cell death [2]. These inclusion bodies are prominent in the postmortem brains of patients who have suffered from these neurodegenerative diseases, and early investigation of inclusion bodies revealed the presence of filamentous a-synuclein (a-syn) [1,2]. a-syn is a small (14.4 kDa) intrinsically disordered protein whose physiological role remains elusive. a-syn has the capability to bind to the SNARE complex and associate with vesicles at the neuronal axon terminus, providing evidence that it may have an impact on neurotransmitter release, vesicle docking, and vesicle trafficking [5–8]. However, upon misfolding, a-syn first forms oligomeric aggregates that eventually undergo fibrilization; these fibrils display the highly ordered cross β-sheets classically found in amyloids [9,10]. These, in turn, form the extended filaments that cause neuropathological changes in the brain and are specifically responsible for PD, LB, and MSA. Diseases caused by a-syn in this manner are called synucleinopathies [11].
The high-resolution structure presented here of filamentous wild-type a-syn is of a helical filament composed of two protofilaments; each turn (or rung) of the filament is comprised of two copies (one per protofilament) of a-syn facing nearly 180° from each other (Figure 1). Between the monomers that make up each protofilament, there is a hydrophobic interface composed of residues 50–57, similar to previously solved structures of filamentous a-syn [12–14]. This interface is stabilized by salt bridges and pseudo-screw symmetry, as previously reported [12,13]. For a-syn, there are seven different missense familial mutations commonly found in patients who have a higher disposition for synucleinopathies (A30P, E46K, H50Q, G51D, A53E, A53T, and A53V) [15–21]. Interestingly, six of these familial mutations lie within the core of the structure and may cause destabilization, resulting in a variety of different fibril morphologies. The presence of polymorphism has been demonstrated particularly well through the analysis of in vitro a-syn fibrils. Fibril twist, crossover distance, packing arrangement, number of protofilaments, interface, and tertiary structure, among others, can vary greatly under different micro- and macro-environments. Many different environmental factors such as pH, salt concentration, temperature, quiescence, and post-translational modifications have an impact on fibril morphology. This has led to the documentation of more than 60 in vitro structural polymorphs of a-syn in the PDB [22,23]. These structural differences in the in vitro filaments can have direct effects on nucleation rates, seeding propensities, and even cytotoxicity [23]. Unfortunately, the ties between these structurally distinct in vitro polymorphs to those found in sarkosyl-insoluble brain-derived structures remain elusive. However, evidence suggests that different polymorphs may influence pathologies [24–26]. This is demonstrated by the difference in a-syn folds of the filaments extracted from patients diagnosed with MSA vs. PD [27].
The formation of the filaments responsible for synucleinopathies is propagated in brain tissue by primary nucleation events in which a-syn monomer spontaneously undergoes structural changes resulting in nucleation. This nucleation site can then recruit additional a-syn monomers to bind, thus elongating the fibril [28,29]. However, there can also be secondary nucleation events in which preformed fibrils are introduced into the cellular environment as “seeds” [30]. These seeding events are significantly more potent at fibril formation and elongation. Remarkably, seeds from a particular polymorph have been shown to recruit wild-type a-syn, provide a structural template, and form filaments expressing the polymorph of the seed regardless of whether the endogenous protein recruited is pathogenic or not [31]. A consequence of this prion-like self-replication is that a-syn fibrils may move from cell to cell, spreading cytotoxic polymorphs.
The introduction of polymorphism has a multifactorial effect on clinical treatments of neurodegenerative diseases. Our understanding of the implications associated with each polymorph on disease progression, pathology, and patient outcomes is very limited. In addition, the differences in folding, packing, or twists of each polymorph introduce complexities in binding sites, affinities, and accessibility for a “one size fits all” drug for synucleinopathies. This is further complicated by evidence that not only are there disease-specific morphisms but each synucleinopathy can exhibit patient-to-patient heterogeneity [32]. Thus, to overcome these challenges, explore new therapeutic targets, understand specific polymorph effects on neuropathology, and develop therapies with patient-specific approaches, solving both patient-derived and in vitro amyloid polymorphs should be explored.
Here, we describe a helical reconstruction workflow that we use to solve the structure of in vitro assembled filamentous a-syn to a global resolution of 2.04 Å. We purify a-syn filaments from a reaction in which fibril seeding material is combined with monomeric a-syn. The fibril seeding material provides a template for fibril elongation via monomer addition over a 6-week incubation period at 37 °C with shaking at 250 rpm. The purified a-syn filaments are then imaged using negative stain transmission electron microscopy (NS-TEM) to evaluate sample integrity and fibril concentration on the grid. The sample is then applied to grids and plunge frozen, and the vitrified grids are used for cryo-EM data collection. We provide a detailed protocol utilizing RELION to reconstruct a high-resolution cryo-EM electron potential map that is then used for building an atomic model of the fibril (Figure 1B, 1C, 1E). The steps presented here may be applied to studies of various amyloid fibrils and accelerate cryo-EM structure determination in the fields of neurodegenerative research and medicine.
Figure 1. Structural features of a-syn fibrils from cryo-EM structures. A. Cryo-EM structure of full-length a-syn fibril depicting two protofilaments (one in red; one in grey). B. Magnified view of a-syn fibril portraying stacked rungs and filament twist. C. Cross-section of a-syn fibril electron potential map displaying two a-syn monomers that comprise each protofilament, approximately 180° from each other. D. Electron potential map of individual β-sheet stacks twisting. E. Model depicting the secondary structure of stacking β-sheets. F. Example of rise measurement for P21 symmetry (red) and C2 symmetry (blue). G. Possible packing symmetry between protofilaments for P21 symmetry (out of register) (red) and C2 symmetry (in register) (blue).
Materials and reagents
Biological materials
1. Plasmid with wild-type a-syn construct inE. coli BL21(DE3)/pET28a-AS [33]
Reagents
1. LB broth (Invitrogen, catalog number: 12780029)
2. Bacto agar (Dot Scientific Inc., catalog number: DSA20030-1000)
3. Magnesium sulfate (MgSO4) (Fisher Scientific, catalog number: 01-337-186)
4. Calcium chloride (CaCl2) (Fisher Scientific, catalog number: BP510-500)
5. Sodium phosphate (NaH2PO4) (Fisher Scientific, catalog number: 01-337-702)
6. Potassium phosphate (KH2PO4) (Fisher Scientific, catalog number: 01-337-803)
7. Sodium chloride (NaCl) (Fisher Scientific, catalog number: S271-500)
8. IPTG (Fisher Scientific, catalog number: BP1755-10)
9. Tris-HCl (Fisher Scientific, catalog number: PRH5125)
10. EDTA (Fisher Scientific, catalog number: AAA1516130)
11. Kanamycin monosulfate (Thermo Scientific, catalog number: J61272.14)
12. SDS-PAGE gels (Bio-Rad, catalog number: 4561096)
13. SDS-PAGE loading dye (Bio-Rad, catalog number: 1610737)
14. Coomassie Brilliant Blue (TCI, catalog number: 6104-59-2)
15. BME vitamins (Sigma-Aldrich, catalog number: B6891-100mL)
16. Sodium azide (Sigma-Aldrich, catalog number: 19-993-1)
17. Studier trace metal mix (Sigma-Aldrich, catalog number: 41106212)
18. Ammonium sulfate (Fisher Scientific, catalog number: A702-500)
19. Deuterium oxide (2H2O) (Cambridge Isotopes Laboratories, catalog number: DLM-4-1L)
20. BioExpress bacterial cell media 10× concentrate (U-13C, 98%; U-15N, 98%; U-D 98%) (Cambridge Isotopes Laboratories, catalog number: CGM-1000-CDN)
21.15N-NH4CI (Cambridge Isotopes Laboratories, catalog number: 39466-62-10)
22.2H-13C-glucose (Cambridge Isotopes Laboratories, catalog number: CDLM-3813-5)
23. Sodium deuteroxide (NaO2H) (Cambridge Isotopes Laboratories, catalog number: DLM-45-100)
24. 2% uranyl acetate (UA) (EMS, catalog number: 22400-2)
25. Ethane (C2H6) (Airgas, catalog number: ET RP35)
26. Liquid nitrogen (LN2) (Airgas, catalog number: NI UHP230LT350)
Solutions
1. Kanamycin stock solution, 1,000× (40 mg/mL) (see Recipes)
2. Kanamycin stock solution, 1,000× (90 mg/mL) (see Recipes)
3. Conditioning plate (see Recipes)
4. Pre-growth media (see Recipes)
5. Lysis buffer (see Recipes)
6. Wash buffer (see Recipes)
7. Growth media (see Recipes)
8. IPTG stock solution (see Recipes)
9. Buffer A (see Recipes)
10. Buffer B (see Recipes)
11. TEN buffer (see Recipes)
12. Saturated ammonium sulfate solution (see Recipes)
13. Fibrilization buffer (see Recipes)
14. 1% uranyl acetate (UA) (see Recipes)
Recipes
1. Kanamycin stock solution, 1,000× (40 mg/mL)
Reagent Final concentration Amount
Kanamycin monosulfate 40 mg/mL 0.4 g
2H2O n/a 10 mL
Total n/a 10 mL
a. Completely dissolve kanamycin monosulfate in 2H2O.
b. Sterilize solution using a 0.22 μm syringe filter and 10 mL syringe.
c. Aliquot 1,000 μL stocks and store at -20 °C until use.
2. Kanamycin stock solution, 1,000× (90 mg/mL)
Reagent Final concentration Amount
Kanamycin monosulfate 90 mg/mL 0.9 g
2H2O n/a 10 mL
Total n/a 10 mL
a. Completely dissolve kanamycin monosulfate in 2H2O.
b. Sterilize solution using a 0.22 μm syringe filter and 10 mL syringe.
c. Aliquot 1,000 μL stocks and store at -20 °C until use.
3. Conditioning plate
Reagent Final concentration Amount
2H2O 70% 700 mL
LB broth 2% 20 g
Bacto agar 1.5% 15 g
H2O n/a Fill to 1,000 mL
Total n/a 1,000 mL
a. Combine reagents in a flask and autoclave at 121 °C, 15 psi for at least 20 min.
b. Allow the media to cool to ~55 °C, then add 1,000 μL of the kanamycin stock solution (1,000×, 40 mg/mL).
c. Pour ~25 mL of media per Petri plate (100 mm) and repeat for the remaining 1 L.
4. Pre-growth media
Reagent Final concentration Amount
2H2O 70% 35 mL
LB broth 2% 1 g
H2O n/a Fill to 50 mL
Total n/a 50 mL
a. Combine reagents in a flask and autoclave at 121 °C, 15 psi for at least 20 min.
b. Allow the media to cool to ~55 °C and then add 50 μL of the kanamycin stock solution (1,000×, 40 mg/mL).
5. Lysis buffer
Reagent Final concentration Amount
NaOH 40 mM 0.80 g
Tris-HCl 20 mM 3.15 g
EDTA 1 mM 0.0585 mg
Triton X-100 0.1% (v/v) 0.5 mL
Total n/a 500 mL
Combine reagents and adjust pH to 8.0 with NaOH.
6. Wash buffer
Reagent Final concentration Amount
NaH2PO4 50 mM 0.34 g
KH2PO4 25 mM 0.17 g
NaCl 10 mM 0.03 g
2H2O n/a Fill to 50 mL
Total n/a 50 mL
a. Combine reagents and adjust the pH to 7.6 with NaO2H.
b. Filter sterilize the solution using a 500 mL filtration system.
7. Growth media
Reagent Final concentration Amount
NaH2PO4 50 mM 6.9 g
KH2PO4 25 mM 3.4 g
NaCl 10 mM 0.58 g
MgSO4 5 mM 1.23 g
CaCl2 0.2 mM 0.03 g
Bacterial cell media 10× 0.1× 10 mL
BME vitamins 100× 0.25× 2.5 mL
Studier trace metals 1,000× 0.25× 0.25 mL
15N-NH4CI 1 g/L 1 g
2H-13C-glucose 8 g/L 8 g
BME vitamins 0.25× 2.5 mL
2H2O n/a Fill to 50 mL
Total n/a 1,000 mL
a. Combine reagents and add 1,000 μL of the kanamycin stock solution (1,000×, 90 mg/mL). Adjust pH to 7.6 with NaO2H.
b. Filter sterilize the solution using a 1,000 mL filtration system.
8. IPTG stock solution
Reagent Final concentration Amount
IPTG 0.5 M 1.2 g
2H2O n/a 10 mL
Total n/a 10 mL
a. Completely dissolve IPTG in 2H2O.
b. Sterilize solution using a 0.22 μm syringe filter and 10 mL syringe.
c. Aliquot 1,000 μL stocks and store at -20 °C until use.
9. Buffer A
Reagent Final concentration Amount
Tris-HCl 30 mM 4.73 g
NaCl 30 mM 1.75 g
H2O n/a 1 L
Total n/a 1 L
a. Dissolve Tris-HCl and NaCl in water while stirring.
b. Adjust pH to 7.4 at 37 °C using 1M NaOH.
10. Buffer B
Reagent Final concentration Amount
Tris-HCl 30 mM 2.36 g
NaCl 1 M 29.22 g
H2O n/a 500 mL
Total n/a 500 mL
a. Dissolve Tris-HCl and NaCl in water while stirring.
b. Adjust pH to 7.4 at 37 °C using 1M NaOH.
11. TEN buffer
Reagent Final concentration Amount
Tris-HCl 30 mM 4.73 g
NaCl 30 mM 1.75 g
EDTA 0.1 mM 29.22 mg
H2O n/a 1 L
Total n/a 1 L
a. Dissolve Tris-HCl, NaCl, and EDTA in water while stirring.
b. Adjust pH to 8.0 at 37 °C using 1 M NaOH.
12. Saturated ammonium sulfate solution
Reagent Final concentration Amount
Ammonium sulfate saturated ~550 g
H2O n/a 1 L
Total n/a 1 L
a. Add ammonium sulfate into water while stirring.
b. Heat gently until all ammonium sulfate is dissolved.
c. Cool to room temperature. Crystals should form to indicate the solution is saturated.
13. Fibrilization buffer
Reagent Final concentration Amount
NaH2PO4 50 mM 1.2 g
EDTA 0.1 mM 5.85 mg
Sodium azide 0.02% 40 µg
H2O 20% 40 mL
2H2O 80% 160 mL
Total n/a 200 mL
a. Add NaH2PO4, EDTA, and 0.02% sodium azide solution into 2H2O and H2O.
b. Adjust pH to 7.4 at 37 °C using 1M NaO2H.
14. 1% uranyl acetate (UA)
Reagent Final concentration Amount
2% uranyl acetate 1% (v/v) 250 μL
H2O n/a 250 μL
Total n/a 500 μL
Mix 1 part of sterile water with 1 part of 2% UA and filter through a Spin-X centrifuge tube with a 0.22 μm filter.
Laboratory supplies
1. 10 mL syringe (BD, catalog number: 309604)
2. 0.22 μm filter (GenClone, catalog number: 25-240)
3. 100 mm × 15 mm Petri dishes (Fisher Scientific, catalog number: S33580A)
4. 500 mL filtration system (Nalgene, catalog number: 595-4520)
5. 1,000 mL filtration system (Fisher Scientific, catalog number: FB12566506)
6. 50 mL conical tubes (VWR, catalog number: 525-1074)
7. 0.45 μm syringe filter (GenClone, catalog number: 25-246)
8. 1.7 mL centrifuge tubes (Denville, catalog number: C2170)
9. Parafilm (Bemis, catalog number: PM996)
10. 0.22 μm Spin-X centrifuge tube filter (Costar, catalog number: 8160)
11. 200 mesh carbon film, copper grids (EMS, catalog number: CF200-CU)
12. Whatman #1 filter paper (Whatman, catalog number: 1001-090)
13. Quantifoil R2/1 200 mesh, copper grids (Quantifoil Micro Tools GmbH, catalog number: Q210CR1)
14. Standard Vitrobot filter paper, Ø 55/20 mm, grade 595 (Ted Pella, catalog number: 47000-100)
Equipment
1. HiTrap Q HP anion exchange column with QFF anion exchange resin (Cytiva, catalog number: 17115401)
2. Stirred cell concentrator (Amicon, catalog number: UFSC05001)
3. Ultracel 3 kDa ultrafiltration disc (Amicon, catalog number: PLBC04310)
4. HiPrep 16/60 Sephacryl S100-HR gel filtration column (Cytiva, catalog number: 17119501)
5. 5424 R microcentrifuge (Eppendorf, catalog number: 05-400-005)
6. 5810 R centrifuge (Eppendorf, catalog number: 022625101)
7. J6-MI high-capacity centrifuge (Beckman Coulter, catalog number: 449598)
8. 1 L centrifuge bottle (Beckman Coulter, catalog number: C31597)
9. myTemp digital incubator (Benchmark Scientific, catalog number: H2200-HC)
10. Pyrex 250 mL Erlenmeyer flask (Corning, catalog number: 4980-250)
11. Floor orbital shaker (Thermo Electron Corporation, catalog number: 19141, model: 480)
12. Denovix UV spectrophotometer/fluorimeter (Denovix, model: DS-11 FX+)
13. Quartz cuvette (Denovix, catalog number: A-70031)
14. ӒKTA pure 25 M (Cytiva, catalog number: 29018226)
15. Grid holder block (Pelco, catalog number: 16820-25)
16. Plasma cleaner (Harrick Plasma Inc., catalog number: PDC-32G)
17. Static dissipator (Mettler Toledo, catalog number: UX-11337-99)
18. Style N5 reverse pressure tweezers (Dumont, catalog number: 0202-N5-PS-1)
19. Talos L120C 120 kV transmission electron microscope (TEM) (Thermo Fisher Scientific or equivalent)
20. Cryo grid box (Sub-Angstrom, catalog number: SB)
21. Vitrobot Mark IV vitrification robot (Thermo Fisher Scientific)
22. Titan Krios G3i 300 kV transmission electron microscope (TEM) (Thermo Fisher Scientific)
23. K3-GIF direct electron detector with energy filter (Gatan Inc., AMETEK)
24. High-performance computing (HPC) cluster with an EPYC Milan 7713P 64-core 2.0 GHz CPU (AMD), 512 GB RAM, 4× RTX A5000 24 GB GDDR6 GPU (NVIDIA), 2× 960 GB Enterprise SSD, mirrored OS, 2× 7.68 TB nVME SSD as 15 TB scratch space, dual-port 25 GbE Ethernet
Software and datasets
1. EPU/AFIS (https://thermofisher.com/smart-epu)
2. SBGrid (https://sbgrid.org/) [34]
3. IMOD (https://bio3d.colorado.edu/imod/) [35]
4. RELION (https://relion.readthedocs.io/en/release-4.0/) [36,37]
5. MotionCor2 (https://emcore.ucsf.edu/ucsf-software) [38]
6. Gctf (https://sbgrid.org/software/titles/gctf) [39]
7. Topaz-filament (https://github.com/3dem/topaz) [40]
8. UCSF ChimeraX (https://www.cgl.ucsf.edu/chimerax/) [41,42]
9. Coot (https://www2.mrc-lmb.cam.ac.uk/personal/pemsley/coot/)
10. PHENIX (https://phenix-online.org/) [43]
Procedure
A. a-synuclein sample preparation
Expression and purification of a-syn protein is performed as reported previously [33]. The protein preparations and fibrilization protocol presented here were developed for joint cryo-EM and NMR studies. Preparations include the use of isotopically labeled reagents that are critical for NMR experiments but are not necessary for cryo-EM. Thus, the a-syn sample preparation protocol may be adapted for cryo-EM only studies by substituting isotopically labeled reagents with a standard equivalent reagent.
A1. a-synuclein protein expression
1. Expression of wild-type a-syn is performed in E. coli BL21(DE3)/pET28a-AS.
2. Plate transformed cells onto the conditioning plate overnight at 37 °C.
3. Inoculate a 50 mL pre-growth flask with a single colony from the overnight conditioning plate and incubate overnight at 220 rpm at 37 °C until OD600 = ~3.
4. Transfer cells into a 50 mL conical tube using aseptic techniques. Centrifuge tubes at 3,200× g for 5 min at 4 °C to form a cell pellet. Decant supernatant and wash with ~20 mL of cold wash buffer.
5. Resuspended cells with the growth media within conical vials. Transfer approximately equal cell quantities into 4× 1 L baffled flasks and fill each to a volume of 250 mL of growth media. Allow cells to grow at 37 °C shaking at 220 rpm until an OD600 of ~1–1.2 is reached. Then, induce a-syn overexpression by adding 1 mL of IPTG stock solution. Incubate at 25 °C with shaking at 200 rpm.
6. After overnight growth, collect cells and combine for harvesting (~15 h post-induction) into a 1 L centrifuge bottle. Centrifuge at 2,500× g for 10 min at 4 °C. Decant the supernatant and wash the cell pellet with the wash buffer to remove residual growth media components.
7. Cell pellets may then be frozen and stored at -80 °C until use.
A2. a-synuclein protein purification
1. Cells may be lysed via heat denaturation, as a-syn is thermostable and will be unaffected. To the conical tubes containing the cell pellet, add 15 mL of lysis buffer and place the conical tubes containing cell paste in boiling water (98 °C) for 30 min. Cool cell lysate on ice. Clear the cell lysate by centrifugation at 3,200× g for 10 min at 4 °C.
2. a-syn should then be precipitated via addition (to 50% v/v) of a saturated ammonium sulfate solution on ice. Collect a-syn precipitate via centrifugation at 100,000× g for 45 min at 4 °C and decant the supernatant, resulting in a fine white precipitate.
3. Equilibrate the HiTrap Q HP anion exchange column with Buffer A using the FPLC system.
4. Resolubilize the a-syn precipitate with ~5 mL of buffer A. Make sure to filter the resolubilized a-syn using a 0.45 μm syringe filter (GenClone). Inject the resolubilized a-syn to bind to the QFF anion exchange resin. Elute using a linear gradient of 0.03–0.6 M NaCl by increasing the proportion of buffer B flow through the column. Collect fractions as they come off the column. In our hands, fractions containing a-syn monomer usually elute at about 0.3 M NaCl.
5. After completion, run SDS-PAGE to check which fractions (gel bands) contain a-syn. Take 20 mL samples from each fraction tube from 20% buffer B to 40% buffer B. Add 20 mL of SDS-PAGE loading dye to each sample tube and heat at 90 °C for 5 min. Run all samples on an SDS-PAGE gel. Use Coomassie Brilliant Blue stain to stain the gel. Examine the stained gel for a-syn overexpression bands. Note that a-syn tends to run at an apparent size of 18 kDa. Pool these fractions.
6. Concentrate the a-syn monomer solution using a stirred cell concentrator using a 3 kDa molecular weight cutoff filter to a final concentration of ~15 mg/mL, measured via UV spectrophotometer and an extinction coefficient of 5,600 M-1·cm-1 at 280 nm. Prewet the concentrator with buffer A before adding a-syn solution to prevent loss of sample to the filter.
7. Equilibrate the 16/60 Sephacryl S-200 HR gel filtration column with TEN buffer with 5× column volume.
8. Inject 1 mL of the concentrated a-syn pool into the loop path of the 16/60 Sephacryl S-100 HR gel filtration column and run the protocol at 0.5 mL/min until the fraction with an apparent mass of 15 kDa at ~97 min.
9. Pool fractions and concentrate to ~15 mg/mL a-syn using a clean stirred cell concentrator and a 3 kDa molecular weight cutoff filter. Prewet the unit and filter with TEN buffer before adding a-syn solution to prevent loss of sample to the filter.
10. Perform standard SDS-PAGE to validate that the monomeric protein is at the expected size of ~17 kDa (Figure 2).
11. Purified a-syn may then be frozen and stored in a -80 °C freezer until use.
Figure 2. SDS-PAGE of purified a-syn. Lane L corresponds to an aliquot of the PageRuler Plus protein ladder. Lane 1 is post–size exclusion chromatography purified a-syn, with an apparent mass of ~17 kDa on an SDS-PAGE gel.
A3. a-synuclein fibrilization
1. Buffer exchange from the TEN buffer to the fibrilization buffer. Add purified a-syn from above to a prewetted (with the fibrilization buffer) stirred cell concentrator and a 3 kDa molecular weight cut-off filter. Dilute 10× with fibrilization buffer and concentrate down to the initial volume. Repeat three times to effectively remove the TEN buffer and completely exchange it to fibrilization buffer.
2. Concentrate purified a-syn protein in the above buffer to 15 mg/mL using 3 kDa cut-off stirred cell concentrators and 0.5 mL aliquot into clean, sterile 1.7 mL Eppendorf tubes.
3. Fibril formation may be seeded with ~50 ng of previously made mature a-syn fibril (in this case, the sample used to determine the PDB ID: 2N0A fibril structure).
Note: To form a fibril without seeding, allow monomeric a-syn to shake at 250 rpm at 37 °C for 6 weeks.
4. Seal the tubes with parafilm for the duration of the incubation.
5. Incubate at 37 °C and shake at 250 rpm continuously for 3 weeks. The viscosity of the fibril solution will greatly increase over time.
6. At the end of 3 weeks, add 100 mL of fibrilization buffer and continue the incubation for 3 weeks under the same conditions.
7. After a total of 6 weeks, the fibrils at a protein concentration of ~13 mg/mL are ready for TEM analysis.
B. Negative stain
Fibrilization can be characterized by thioflavin-T (ThT) assays, which leverage the fluorescence signal observed when thioflavin-T binds to fibrils, a property not observed in the presence of purified protein monomers [44]. Although this method is powerful and can even detail fibrilization kinetics, there are limitations in the technique. Specifically, this assay cannot specify whether fibrils are twisting, if they span several micrometers in length, or if they are small fragments tens of nanometers in length. For high-resolution cryo-EM structure determination, fibrils should be both twisting and span several crossovers; if fibrils have one or less crossovers, they are too short for cryo-EM data collection. The crossover occurs when a single protofilament turns 180° around the fibril axis, a feature that can be observed by negative stain transmission electron microscopy (NS-TEM) (Figures 3 and 4). Additionally, fibrils should be concentrated to a point where several fibrils span the micrograph but are not crowded or overlapping. This ensures there are enough individual particles for the reconstruction process. In vitro preparations should consist mostly of fibril samples. If excessive aggregation is observed, then it is best to dilute or centrifuge the sample to remove these large aggregates. This is important as consistently having material that is too large will change the thickness of the ice and yield lower-quality cryo-EM data. Samples extracted from tissue may have significant background and tissue-specific material that was not removed during extraction. Since these samples are precious and additional purification may not be possible, fibrils that can be clearly differentiated from the background and are in sufficient quantities may move forward in the workflow, though a larger dataset may be needed for reconstruction. To determine if the fibrils possess these qualities, we perform NS-TEM with the following procedure to test a range of sample concentrations. We found that a concentration of 6.5 mg/mL (i.e., 1:1 ratio of sample to buffer) was best for our in vitro sample on the grid.
1. Place the desired number of 200 mesh carbon film, copper EM grids on a grid holder block and use a plasma cleaner (PDC-32G or equivalent system) to glow discharge grids under a 100-micron vacuum for 30 s on low (Figure 3, step 1).
2. Cut a piece of parafilm to approximately 2” × 4” and demagnetize with a static dissipater (Figure 3, step 2).
3. Retrieve one glow-discharged EM grid using style N5 reverse pressure tweezers or similar tweezers (Figure 3, step 3).
4. Spot two 50 μL drops of sterile, Nanopure water and two 50 μL drops of 1% UA onto the piece of parafilm. Ensure the drops do not touch (Figure 3, step 4).
5. Apply 4 μL of the sample to the EM grid and allow the sample to incubate at room temperature for 1 min (Figure 3, step 5).
6. Blot away the liquid by touching the edge of the EM grid to a piece of filter paper (Figure 3, step 6).
7. Wash the EM grid by touching the face of the EM grid to the first drop of water and then blot away the liquid as in step B6. Repeat, but this time wash with the second drop of water (Figure 3, step 7).
8. Pre-stain the EM grid by touching the face of the EM grid to the first drop of 1% UA, then blot away the stain as in step B6 (Figure 3, step 8).
9. Stain the grid by holding the face of the EM grid to the second drop of 1% UA for 15 s, then blot away the stain as in step B6 (Figure 3, step 9).
10. Allow the EM grid to dry for at least 5 min at room temperature before storing the grid in a grid box (Figure 3, step 10). Store the grid box in a desiccator or humidity-controlled room until imaging.
Figure 3. Negative stain protocol. Detailed steps for preparing negative stain grids of a-syn fibrils. The protocol yields lightly stained fibrils, allowing for the visualization of twisting fibrils comprised of two protofilaments (Figure 4). The procedure is repeated, spanning a range of fibril concentrations that are imaged by transmission electron microscopy.
Figure 4. Negative stain TEM analysis of a-synuclein fibrils. Representative micrograph of fibrils lightly stained with 1% UA. The in vitro fibrils are comprised of two protofilaments (arrows) and appear to be twisting with distinct crossover points (stars). These fibrils are long enough to span the micrograph, demonstrate an optimized preparation, are not overcrowded indicating appropriate concentration, and have minimal overlap to allow for adequate particle picking. Additionally, the micrograph did not appear to contain aggregates, contaminates, or extra biological material. This is optimal for data collection/processing and represents an encouraging negative stain screen. Scale bar, 100 nm.
11. Repeat for additional sample dilutions to assess the sample conditions that may be best suited for cryo-EM analysis. We imaged the sample at a concentration of 13 mg/mL (undiluted), 6.5 mg/mL (2× dilution), and 2.6 mg/mL (5× dilution). We found that a concentration of 6.5 mg/mL showed the best sample distribution on the grid (Figure 4).
Note: Since fibrilization conditions greatly impact the length of the fibrils and thus the sample distribution on the grid, it is important to test each sample by NS-TEM before sample vitrification and cryo-EM data collection.
12. Image grids on a Talos L120C 120 kV TEM or equivalent microscope at a pixel size of 1.58 Å and a total electron dose of ~25 e-/Å2.
C. Sample vitrification
Basic sample vitrification for single particle analysis has become routine in the cryo-EM field. Here, we present a brief workflow of the vitrification process using the Vitrobot Mark IV with blotting conditions that yield grids suitable for cryo-EM data collection.
1. Using a plastic syringe, add 60 mL of distilled water to the Vitrobot Mark IV water reservoir.
2. Turn on the Vitrobot Mark IV and set the chamber temperature to 20 °C and the relative humidity to 95%.
3. Attach standard Vitrobot filter paper to the blotting pads and allow the system to equilibrate to the conditions set in step C2 (~15 min).
4. Using a plasma cleaner or equivalent system, glow-discharge R2/1 200 mesh, copper grids.
5. Use liquid nitrogen (LN2) to cool the Vitrobot foam dewar, ethane cup, and metal spider.
6. Once the setup has cooled, condense the ethane in the ethane cup. Be sure to monitor ethane and LN2 levels throughout the vitrification process.
7. On the Vitrobot, set the wait time to 60 s and set the drain time to 0.5 s. For blot force and blot time, it is usually necessary to test a range of parameters that work best. For these fibrils, a blot time between 4 and 5 s and a blot force of -1 to +2 worked well.
Note: There is variance between Vitrobots; thus, optimization of blotting force and blot time may be necessary for the specific equipment being used.
8. Using the Vitrobot tweezers, pick up a grid and attach the tweezers to the Vitrobot. Selectcontinue on the screen to raise the tweezers and mount the foam dewar in place. Follow the prompts on the screen to bring the tweezers and dewar into position for sample application.
9. Apply 4 mL of the fibrils to the carbon side of the grid. Selectcontinue to begin the wait time; then, the system will automatically blot and plunge the sample into liquid ethane.
10. Once the system has plunged the specimen into the cryogen, transfer the vitrified grid to a labeled grid box and store appropriately.
Note: A complete guide on plunge freezing using the Vitrobot can be found in unit 2 of the Getting Started with Cryo-EM videos (https://youtube.com/playlist?list=PL8_xPU5epJdfd5fM2CjQItR-iRlIEIJk8&si=NAqencnr2wpuZg9B).
11. Repeat steps C7–9 for any additional grids. In addition to duplicate grids, it is always beneficial to test a range of blotting conditions and/or sample concentrations. Cryo-EM data was collected on a grid with a blot time of 4 s and a blot force of +2 at a protein concentration of ~6.5 mg/mL.
D. Cryo-EM data collection
Data collection parameters should be tailored to the resources available, and thus users should work closely with EM facility staff to optimize the data collection parameters for their individual sample. Here, the data was acquired on a Titan Krios G3i FEG-TEM. The microscope is operated at 300 kV and is equipped with a Gatan K3 direct electron detector and a BioQuantum energy filter set at 20 eV. Correlated double sampling (CDS) was used to collect dose-fractionated micrographs using a defocus range of -0.5 to -2.5 μm with increments of 0.25 μm. Micrographs were collected at a magnification of 105,000× with a pixel size of 0.834 Å and a total dose of 40 e-/Å2 (1 e-/Å2/frame). On average, ~250 movies were collected per hour using EPU/AFIS (Thermo Fisher Scientific) acquiring three shots per hole and multiple holes per stage movement. A representative micrograph at an estimated defocus of -2.0 μm shows twisting fibrils suspended in vitreous ice (Figure 5).
Figure 5. Representative cryo-EM micrographs of a-synuclein fibrils. Motion-corrected micrograph of vitrified a-syn fibrils at an estimated defocus of -2.0 μm. The fibrils are comprised of two protofilaments (arrows) that are twisting at distinct crossover points (stars). Twisting fibrils are critical for high-resolution structure determination. Scale bar, 100 nm.
E. Cryo-EM data processing of a-synuclein fibrils
Cryo-EM structure determination of amyloid fibrils has revolutionized the fields of neuroscience and neurodegenerative medicine, providing key structural details that were previously unattainable by other methods. Here, we provide a data processing protocol that is both detailed and reproducible to serve as a starting point for those new to cryo-EM and helical reconstruction workflows. The raw micrographs, gain file, and the detector mtf file can be accessed at EMPIAR-12229, allowing users to work through the steps below before applying the workflow to new experimental data. We must note that all data sets are unique and possess their own challenges, but this workflow should greatly improve the user’s ability to resolve amyloid fibril structures. Finally, as with any software, it is best to first become accustomed to the program by completing the appropriate tutorial datasets. We highly encourage readers to first complete the RELION single particle tutorial (https://relion.readthedocs.io/en/release-4.0/SPA_tutorial/index.html) before proceeding with the steps below [36].
Creating a RELION Project
Create a directory that will house the entire RELION project. For simplicity, call this directory a-syn_data_processing. Within this directory, you should have two files titled gain.mrc andk3-CDS-300keV-mtf.star and a subdirectory titled Micrographs that contains all the raw movie frames in tiff format. These files can be downloaded from EMPIAR-12229. Now that your directories are organized, cd to thea-syn_data_processing directory, this will serve as the RELION parent directory for all subsequent jobs. Launch RELION by running relion & in the terminal. The “&” will allow RELION to run in the background in case the terminal is needed for additional commands. As a final note, we have listed the input files for each job based on our RELION project so there will be discrepancies in job numbers between our project and yours. Thus, it is important to use the proper input path file for your project at each step. For each step, we have detailed where the input file comes from (i.e., the step the file was generated in) to ensure successful reconstruction of the EMPIAR-12229 dataset.
Allocating computational resources when running RELION jobs
RELION uses a Compute and Running tab to allocate computational resources based on user-defined parameters. These parameters are completely dependent on the resources available to each individual. Thus, rather than detailing these parameters for each step, here we have outlined the Compute and Running parameters that work well for our HPC cluster with slurm queueing system. However, these parameters may not work for your computational setup, and you may need to seek the advice of IT professionals at your institute. We have also included the Compute and Running tabs in Video 1 for each job as an additional resource for determining these parameters.
Video 1. RELION-4 parameters for the helical reconstruction of a-synuclein fibrils. Screenshots of the RELION GUI for each step from section E are provided for easier visualization of input parameters. Additional information regarding the origin of the input files is provided, as the file names will vary from project to project depending on the number of RELION jobs run.
Compute
Use parallel disk I/O? Yes
Number of pooled particles: 30
Skip padding? No
Pre-read all particles into RAM? No
Copy particles to scratch directory: Leave Blank
Combine iterations through disc? No
Use GPU acceleration? Yes
Which GPUs to use: Leave Blank
Running (GPU jobs):
Number of MPI procs: 5
Number of threads: 6
Submit to queue? Yes
Queue name: a5000
Queue submit command: sbatch
Standard submission script: ../../../../../../share/sbatch/relion_template_gpu.sh
Minimum dedicated cores per node: 1
Additional arguments: Leave Blank
Running (CPU jobs):
Number of MPI procs: 20
Submit to queue? Yes
Queue name: cpu
Queue submit command: sbatch
Standard submission script: ../../../../../../share/sbatch/relion_template_cpu.sh
Minimum dedicated cores per node: 1
Additional arguments: Leave Blank
In addition to the protocol below, a workflow diagram and a video of the RELION GUI with parameters for each step are provided (Figure 6, Video 1).
Figure 6. Helical reconstruction workflow for a-synuclein fibrils using RELION. Overview of each RELION job utilized to reconstruct a-syn fibrils to ~2.0 Å. Each job corresponds to the step number in section E and to those in Video 1.
1. Import
First, import the raw movie frames into RELION for data processing. Select the Import job, ensure Raw input files is set to the location of the movie frames, use the “*” argument to select all the tiff files in the directory, set the additional parameters below, and click the Run! button.
Movies/mics:
Import raw movies/micrographs: Yes
Raw input files: Micrographs/*.tiff
Optics group name: opticsGroup1
MTF of the detector: k3-CDS-300keV-mtf.star
Pixel size (Angstrom): 0.834
Voltage (kV): 300
Spherical aberration (mm): 2.7
Amplitude contrast: 0.1
Beamtilt in X (mrad): 0
Beamtilt in Y (mrad): 0
Others:
Import other node types? No
The output log will display 5,193 micrographs imported.
2. Motion correction
The raw movie frames from the previous job (movies.star) must now be aligned. The data was collected with a dose per frame of 1 e-/Å2 over 40 frames for a total dose of 40 e-/Å2. Note that the EER fractionation parameter will be ignored by RELION since these images were collected on a Gatan K3 detector and are tiff files. Perform motion correction by using the MotionCor2 program [38]. Tell RELION where the program is located via the MOTIONCOR2 executable parameter. Your computational setup will be different, and MotionCor2 may be saved in a different location, so the executable path may be different. In the terminal, run which motioncor2 to determine the correct path for the program. Similarly, your computational setup will dictate the number of GPUs available. Our setup includes multiple nodes, and each can run 4 GPUs concurrently. In the RELION GUI, use Which GPUs to use to indicate the GPUs available for your setup; leaving this blank will automatically allocate the GPUs. Select the Motion correction job, set Input movies STAR file to themovie.star file from step 1, set the following parameters and update any paths or parameters that are specific to your computational setup, and then click theRun! button.
I/O:
Input movies STAR file: Import/job001/movies.star
First frame for corrected sum: 1
Last frame for corrected sum: -1
Dose per frame (e-/Å2): 1
Pre-exposure (e-/Å2): 0
EER fractionation: 32
Write output in float16? Yes
Do dose-weighting? Yes
Save non-dose weighted as well? No
Save sum of power spectra? Yes
Sum power spectra every (e-/Å2): 4
Motion:
Bfactor: 150
Number of patches X, Y: 5, 5
Group frames: 1
Binning factor: 1
Gain-reference image: gain.mrc
Gain rotation: 180 degrees (2)
Gain flip: Flip left to right (2)
Defect file: Leave blank
Use RELION’s own implementation? No
MOTIONCOR2 executable: /programs/x86_64-linux/motioncor2/1.3.1/motioncor2
Which GPUs to use: 0,1,2,3
Other MOTIONCOR2 arguments: Leave blank
This job will take several hours to run and will generate a corrected_micrographs.star file. If interested, you may open the logfile.pdf to visualize the results from the job. Under the Finished Jobs list click on the Motion correction job. This will update the Current: job display (located in the center of the GUI) to your Motion correction job and upload the results to the user interface. On the right side of the RELION GUI there is a drop-down menu called Display: that allows the user to visualize outputs from the finished job. Click on the drop-down menu and select Out: logfile.pdf. A new window will appear with the results of the job. Subsequent logfile.pdf files from finished jobs can be opened this way.
3. CTF estimation
Now, estimate the CTF values for the motion-corrected micrographs from the previous step; these are stored in the corrected_micrographs.star file. Use Gctf to estimate CTF values [39]. In the terminal, run which Gctf to determine the correct executable path for your setup. In the RELION GUI, select the CTF estimation job, set Input micrographs STAR file to the corrected_micrographs.star file from step 2, set the following parameters and update any paths specific to your setup, and then click the Run! button.
I/O:
Input micrographs STAR file: MotionCorr/job002/corrected_micrographs.star
Use micrograph without dose-weighting? No
Estimate phase shifts? No
Amount of astigmatism (Å): 100
CTFFIND-4.1:
Use CTFFIND-4.1? No
FFT box size (pix): 512
Minimum resolution (Å): 30
Maximum resolution (Å): 5
Minimum defocus value (Å): 5000
Maximum defocus value (Å): 50000
Defocus step size (Å): 500
Gctf:
Use Gctf instead? Yes
Gctf executable: /programs/x86_64-linux/gctf/1.06/bin/Gctf
Ignore ‘Searches’ parameters? Yes
Perform equi-phase averaging? Yes
Other Gctf options: Leave blank
Which GPUs to use: 0,1,2,3
This job results in a micrographs_ctf.star file and a logfile.pdf file. The logfile.pdf contains a graphical representation of the metadata related to micrograph defocus, astigmatism, max resolution, and figure of merit values. These values will be used in the upcoming steps to filter the micrograph dataset.
4. Subset selection (defocus filter)
Extensive testing has shown that using stringent parameters during the micrograph curation steps allows for a segment picking neural network that performs better than one trained on the entire data set. The following steps will use CTF estimation results to curate a set of micrographs for manual picking. Those picks will then be used to train the Topaz neural network [40]. Finally, a modified version of Topaz called Topaz-filament, which allows for picking filamentous structures, is optimized on a small subset of micrographs before applying the neural network to our entire dataset [40].
Open the logfile.pdf from the CTF estimation job and use the values provided in this file to eliminate any outliers or suboptimal micrographs. Filter the dataset based on defocus, astigmatism, max resolution, and figure of merit values using a series of Subset Selection jobs. Select the Subset Selection job, input the following parameters and update OR select from micrograph.star to the micrographs_ctf.star file from step 3, and then click the Run! button.
I/O:
Select classes from job: Leave blank
OR select from micrograph.star: CtfFind/job003/micrographs_ctf.star
OR select from particles.star: Leave blank
Class options:
Automatically select 2D classes? No
Re-center the class averages? No
Regroup the particles? No
Subsets:
Select based on metadata value? Yes
Metadata label for subset selection: rlnDefocusU
Minimum metadata value: -9999
Maximum metadata value: 25000
OR: select on image statistics? No
OR: split into subsets? No
Duplicates:
OR: remove duplicates? No
This job reduces the number of micrographs from 5,193 to 4,858 based on a maximum defocus value of 2.5 mm (25,000 Å).
5. Subset selection (astigmatism filter)
Filter the micrograph subset from step 4 by the astigmatism values in the CTF estimation logfile.pdf file. Select the Subset Selection job type, set OR select from micrograph.star to the micrographs.star file from step 4, input the following parameters, and then click the Run! button.
I/O:
Select classes from job: Leave blank
OR select from micrograph.star: Select/job004/micrographs.star
OR select from particles.star: Leave blank
Class options:
Automatically select 2D classes? No
Re-center the class averages? No
Regroup the particles? No
Subsets:
Select based on metadata value? Yes
Metadata label for subset selection: rlnCtfAstigmatism
Minimum metadata value: -9999
Maximum metadata value: 700
OR: select on image statistics? No
OR: split into subsets? No
Duplicates:
OR: remove duplicates? No
This job reduces the number of micrographs from 4,858 to 3,390 micrographs.
6. Subset selection (max resolution filter)
Further filter the micrograph subset from step 5 by the max resolution values from the CTF estimation logfile.pdf file. Select the Subset Selection job, set OR select from micrograph.star to the micrograph.star file from step 5, set the following parameters, and click the Run! button.
I/O:
Select classes from job: Leave blank
OR select from micrograph.star: Select/job020/micrographs.star
OR select from particles.star: Leave blank
Class options:
Automatically select 2D classes? No
Re-center the class averages? No
Regroup the particles? No
Subsets:
Select based on metadata value? Yes
Metadata label for subset selection: rlnCtfMaxResolution
Minimum metadata value: -9999
Maximum metadata value: 4
OR: select on image statistics? No
OR: split into subsets? No
Duplicates:
OR: remove duplicates? No
This job reduces the number of micrographs from 3,390 to 910 micrographs.
7. Subset selection (figure of merit filter)
Lastly, filter the micrograph subset from step 6 by the figure of merit values from the CTF estimation logfile.pdf file. Select the Subset Selection job, set OR select from micrograph.star to the micrograph.star file from step 6, set the following parameters, and then click the Run! button.
I/O:
Select classes from job: Leave blank
OR select from micrograph.star: Select/job021/micrographs.star
OR select from particles.star: Leave blank
Class options:
Automatically select 2D classes? No
Re-center the class averages? No
Regroup the particles? No
Subsets:
Select based on metadata value? Yes
Metadata label for subset selection: rlnCtfFigureOfMerit
Minimum metadata value: 0.065
Maximum metadata value: 0.9
OR: select on image statistics? No
OR: split into subsets? No
Duplicates:
OR: remove duplicates? No
This job reduces the number of micrographs from 910 to 774 micrographs.
8. Subset selection (2 sets of 20 micrograph)
From the remaining 774 micrographs, generate 2 sets of 20 micrographs. The first set of micrographs will be used for manual picking and training the neural network. The second set of 20 micrographs will be used to test and optimize the picking thresholds that will then be applied to the entire dataset. Select the Subset Selection job, set OR select from micrograph.star to the micrographs.star file from step 7, and then click the Run! button.
I/O:
Select classes from job: Leave blank
OR select from micrograph.star: Select/job022/micrographs.star
OR select from particles.star: Leave blank
Class options:
Automatically select 2D classes? No
Re-center the class averages? No
Regroup the particles? No
Subsets:
Select based on metadata value? No
OR: select on image statistics? No
OR: split into subsets? Yes
Randomise order before making subset? No
Subset size: 20
OR: number of subsets: 2
Duplicates:
OR: remove duplicates? No
This step results in two STAR files labeled micrographs_split1.star and micrographs_split2.star. Each star file contains 20 micrographs.
9. Manual picking
Select the Manual picking job, set Input micrographs to the micrographs_split1.star file from step 8, set the additional parameters below, and then click the Run! button. A new window will appear with 20 rows (one per micrograph) with the micrograph name, apick button, the number of picks, a CTF button, and the defocus estimate for that micrograph. Click on the pick button to launch a new window for the specified micrograph. Use the left mouse button and click at one end of a fibril, and then click a second time at the opposite end of the fibril. This creates a line segment between the two endpoints defined by the user. The segments will be used for the particle extraction job in subsequent steps. Repeat this process until all the fibrils are picked. Ensure segments do not overlap; if fibrils contain curvature, increase the number of segments that make up the filament (Figure 7A). When done picking, right-click on the micrograph and select Save STAR with coordinates, close the micrograph, and repeat the process for the remaining 19 micrographs. If you need to remove points, use the center button and click over an existing point to remove it. Ensure that all the micrographs have an even number of picks (i.e., one start point and one end point per segment) and that segments are centered over fibrils. When done picking from all 20 micrographs, close the window to finalize the job.
Figure 7. Manual picking, 2D class selection, and auto-picking threshold determination. A. Micrograph with examples of manually picked segments (step 9). Each “end” of the segment is selected by the user (indicated by the stars). The endpoints are then linked by a line (indicated by an arrow); this region will be divided into particles based on the user-defined interbox distance. Each new color represents a new segment that has been manually picked. B. Schematic of interbox distances (step 10). The filament that is shown is a region that has been selected for particle picking. RELION will use a user-defined “box” to select as a particle. The interbox distance shown is the distance in which no overlap from previous boxes is present (i.e., the region that is unique to each box). C. 2D classes from manually picked particles (step 11). The green boxes indicate the classes selected to use for neural network training (step 12). D. Micrographs depicting trained neural network auto-picking results from different threshold values. As the threshold for picking is decreased, the stringency in which the neural network determines whether the feature fits the trained model is decreased, initially resulting in an increase in picked particles. However, as the threshold continues to decrease, the neural network starts to categorize “noise” as pickable particles.
I/O:
Input micrographs: Select/job023/micrographs_split1.star
Pick start-end coordinates helices? Yes
Use autopick FOM threshold? No
Display:
Particle diameter (Å): 100
Scale for micrographs: 0.2
Sigma contrast: 3
White value: 0
Black value: 0
Lowpass filter (Å): 20
Highpass filter (Å): -1
Pixel size (Å): -1
OR: use Topaz denoising? No
Colors:
Blue<>red color particles? No
The output log will list the total number of picks (start and end points). Here, we picked 414 particles (i.e., 207 segments) from 20 micrographs, and the coordinates are saved to themanualpick.star file located in the directory for this job. The total number of segments may vary due to differences in picking, but ensure picks are made on all 20 micrographs.
Note: The parameters in the Display tab are for visualization purposes only and do not impact downstream processing steps.
Note: We observed that in some versions of RELION there is a bug that results in an empty coordinate file from the Manual picking job. To bypass this error, simply select the Manual picking job from the Finished jobs section and then click on the Continue! button. This will reopen the manual picking GUI. Then, close the window; the coordinate file should now be updated with all the picks saved. There is no need to repick particles or change any settings.
10. Particle extraction (manual picks)
The manually picked segments must now be processed to extract particles for 2D classification. In principle, this step will take user-defined parameters to then cut the segments into individual particles for downstream steps (Figure 7B). This is achieved by providing the number of unique asymmetrical units and helical rise (Å) values in the helix tab. RELION will use these values to establish an interbox distance, i.e., the spacing between each particle, that will separate overlapping 360-pixel boxes that traverse the length of the segment (Figure 7B). Here, we have set the interbox distance to ~38.5 Å (4.82 Å × 8) to increase the number of particles for training purposes. This value will be expanded later once auto-picking is complete. Select the Particle extraction job, set Micrograph STAR file to the micrograph_split1.star file from step 8, set Input coordinates to the manualpick.star file from step 9, and then click the Run! button.
I/O:
Micrograph STAR file: Select/job023/micrographs_split1.star
Input coordinates: ManualPick/job024/manualpick.star
OR re-extract refined particles? No
OR re-center refined coordinates? No
Write output in float16? Yes
Extract:
Particle box size (pix): 360
Invert contrast? Yes
Normalize particles? Yes
Diameter background circle (pix): -1
Stddev for white dust removal: -1
Stddev for black dust removal: -1
Rescale particles? No
Use autopick FOM threshold? No
Helix:
Extract helical segments? Yes
Tube diameter (Å): 140
Use bimodal angular priors? Yes
Coordinates are start-end only? Yes
Cut helical tubes into segments? Yes
Number of unique asymmetrical units: 8
Helical rise (Å): 4.82
This job resulted in 5,919 particles extracted to a pixel size of 0.834 Å/pix with a box size of 360 pixels. Differences in particle counts are due to differences in the number of segments picked during the manual picking step. Aim for at least 4,000 particles at this stage.
Note: Amyloid structures have a consistent helical rise of ~4.8 Å. This estimate is sufficient for this stage of processing, as the helical rise will be optimized in subsequent steps.
11. 2D classification (manual picks)
Although the particles were manually picked and thus should be free from suboptimal picks or background noise, we prefer to perform a round of 2D classification to curate the particles that will be used to train the Topaz neural network. Select the 2D classification job, set Input images STAR file to the particles.star file from step 10, set the parameters below, and then click on the Run! button.
I/O:
Input images STAR file: Extract/job029/particles.star
CTF:
Do CTF-correction? Yes
Ignore CTFs until first peak? No
Optimisation:
Number of classes: 20
Regularisation parameter T: 2
Use EM algorithm? Yes
Number of EM iterations: 20
Use VDAM algorithm? No
Mask diameter (Å): 285
Mask individual particles with zeros? Yes
Limit resolution E-step to (Å): 10
Center class averages? Yes
Sampling:
Perform image alignment? Yes
In-plane angular sampling: 2
Offset search range (pix): 5
Offset search step (pix): 1
Allow coarser sampling? No
Helix:
Classify 2D helical segments? Yes
Tube diameter (Å): 140
Do bimodal angular searches? Yes
Angular search range-psi (deg): 6
Restrict helical offsets to rise: Yes
Helical rise (Å): 4.82
Due to the small number of particles and the small number of classes, this job should only take a couple of minutes to run. The final classes can be visualized by clicking on the Display: drop-down menu and selectingout: run_it020_optimiser.star. A RELION display GUI will appear; check the box next to Sort images on: and select rlnClassDistribution from the drop-down menu, then click Display! to see the classes sorted with the most populated classes at the top (Figure 7C). Close the window when done.
12. Subset selection (2D classes for Topaz training)
Next, use the Subset selection job to select the best classes to train the Topaz neural network. Set Select classes from job to the run_it020_optimiser.star file from step 11, set the additional parameters below, and click the Run! button. This will launch a RELION display GUI. Check the box next to Sort images on: and select rlnClassDisribution, then click the Display! button. This will look identical to the previous step where we visualized the classes, but now you use the left mouse button to select all the classes to move to the next step (Figure 7C). Once done, right-click and select Save selected classes, then close the display window.
I/O:
Select classes from job: Class2D/job030/run_it020_optimiser.star
OR select from micrograph.star: Leave blank
OR select from particles.star: Leave blank
Class options:
Automatically select 2D classes? No
Re-center the class averages? Yes
Regroup the particles? No
Subsets:
Select based on metadata values? No
OR: select on image statistics? No
OR: split into subsets? No
Duplicates:
OR: remove duplicates? No
This job resulted in 15 classes selected with 5,712 particles (Figure 7C, green boxes). Your values may be slightly different at this step due to differences in manual picking, but the key is to select classes that appear fibrillar in nature (Figure 7C, green boxes).
13. Auto-picking (Topaz training)
Use the curated particle stack to train a new Topaz neural network. It is critical that the executable path within the Topaz tab directs RELION to the topaz-filament program [40]. The path here is to where topaz-filament is located on our HPC cluster, but this may be different for your setup. If you are unsure where this program is located, you may attempt to locate the program path by running the which topaz-filament command from the terminal. Select the Auto-picking job, set Input micrographs for autopick to the micrographs_selected.star file from step 9, in the Topaz tab set Particles STAR file for training to the particles.star file from step 12, set the additional parameters below and modify the executable path to fit your computational setup, and then click on the Run! button.
I/O:
Input micrographs for autopick: ManualPick/job024/micrographs_selected.star
Pixel size in micrographs (Å): -1
Use reference-based template-matching? No
OR: use Laplacian-of-Gaussian? No
OR: use Topaz? Yes
Laplacian:
This tab is ignored since we opted to use Topaz in the I/O tab.
Topaz:
Topaz executable: /programs/x86_64-linux/system/sbgrid_bin/topaz-filament
Particle diameter (Å): 140
Perform topaz picking? No
Perform topaz training? Yes
Nr of particle per micrograph: 300
Input picked coordinates for training: Leave blank
OR train on a set of particles? Yes
Particles STAR file for training: Select/job032/particles.star
Additional topaz arguments: Leave blank
References:
This tab is ignored since we opted to use Topaz in the I/O tab.
Autopicking:
Use GPU acceleration? Yes
All other parameters on this tab are ignored since we opted to use Topaz in the I/O tab.
Helix:
This tab is ignored since we opted to use Topaz in the I/O tab.
This job results in a trained Topaz model titled model_epoch10.sav which is saved in the folder for this job.
Note: Topaz training is not parallelized, so the job will only use one MPI process.
14. Auto-picking (Topaz picking optimization)
The trained Topaz model will be applied to a subset of 20 micrographs to test how the model performs before it is applied to the entire dataset. For topaz-filament to pick segments and not individual particles as in traditional single particle analysis, the additional flags for filament (-f) and threshold (-t) must be provided in the Additional topaz arguments box. Additionally, an integer value must be provided after the threshold flag. This threshold determines how many particles are picked. A lower threshold results in more particles, but if the threshold is too low, then the model will start picking noise. With any new trained Topaz neural network, we test a range of threshold values, typically from -6 to 0, to see which threshold works best (Figure 7D). Each threshold value will be its own job. Select the Auto-picking job, set Input micrographs for autopick to the micrographs_split2 from job 8, in the Topaz tab set Trained topaz model to the model_epoch10.sav file from step 13, set the parameters below, and click the Run! button. To test additional thresholds, once the first Auto-picking job is complete, click on the job in the Finished jobs list and then click on the Auto-picking job to load the previous settings. Now, simply change the threshold value in the Additional topaz arguments and click the Run! button. Repeat this process for any threshold that you would like to test. We tested thresholds -6, -5, -4, -3, -2, -1, and 0, and found that threshold -5 worked best for the dataset (Figure 7D). We have also included an extreme case with a threshold of -10 to better visualize bad picks that would be unsuitable for further processing (Figure 7D).
I/O:
Input micrographs for autopick: Select/job023/micrographs_split2.star
Pixel size in micrographs (Å): -1
Use reference-based template-matching? No
OR: use Laplacian-of-Gaussian? No
OR: use Topaz? Yes
Laplacian:
This tab is ignored since we opted to use Topaz in the I/O tab.
Topaz:
Topaz executable: /programs/x86_64-linux/system/sbgrid_bin/topaz-filament
Particle diameter (Å): 140
Perform topaz picking? Yes
Trained topaz model: AutoPick/job033/model_epoch10.sav
Perform topaz training? No
Additional topaz arguments: -f -t -5
References:
This tab is ignored since we opted to use Topaz in the I/O tab.
Autopicking:
Use GPU acceleration? Yes
All other parameters on this tab are ignored since we opted to use Topaz in the I/O tab.
Helix:
This tab is ignored since we opted to use Topaz in the I/O tab.
A picking threshold of -5 resulted in 688 segments (1,376 particles, i.e., endpoints) from 20 micrographs.
Note: Topaz picking is parallelized so multiple MPI processes can be run simultaneously; we typically run 20 MPI processes for this job. This setting can be found in the Runningtab and is dependent on the computational resources available.
15. Auto-picking (Topaz picking on the entire dataset)
The trained Topaz model and the optimized picking threshold are now applied to the entire dataset to select segments for downstream processing. As detailed previously, upload the settings from the best picking job (threshold -5), update Input micrographs for autopick to the micrographs.star file from step 4, and click the Run! button.
I/O:
Input micrographs for autopick: Select/job004/micrographs.star
Pixel size in micrographs (Å): -1
Use reference-based template-matching? No
OR: use Laplacian-of-Gaussian? No
OR: use Topaz? Yes
Laplacian:
This tab is ignored since we opted to use Topaz in the I/O tab.
Topaz:
Topaz executable: /programs/x86_64-linux/system/sbgrid_bin/topaz-filament
Particle diameter (Å): 140
Perform topaz picking? Yes
Trained topaz model: AutoPick/job033/model_epoch10.sav
Perform topaz training? No
Additional topaz arguments: -f -t -5
References:
This tab is ignored since we opted to use Topaz in the I/O tab.
Autopicking:
Use GPU acceleration? Yes
All other parameters on this tab are ignored since we opted to use Topaz in the I/O tab.
Helix:
This tab is ignored since we opted to use Topaz in the I/O tab.
This job results in 156,526 segments (313,052 particles, i.e., endpoints) from 4,858 micrographs.
16. Particle extraction (large box size)
For helical reconstruction methods, the helical twist and rise values are critical for cryo-EM data processing. The helical twist can be estimated from 2D class averages with large box sizes that span the fibril crossover distance (Figure 8A, 8B, 8D). Here, extract the particles to a box size of 864 pixels (~720 Å) so we can estimate the crossover distance in subsequent steps. At this stage in the processing, there is no need for high-resolution information, so the box size is rescaled to 144 pixels (i.e., binning to a pixel size of 5.004 Å/pixel). Alternatively, users may estimate the crossover distance from cryo-EM micrographs (typically those with higher defocus values are easier to visualize) or from negative stain TEM micrographs. However, extraction at a larger box size is still necessary to generate an initial reference for 3D reconstruction. Select the Particle extraction job, set Micrograph STAR file to the micrographs.star file from step 4, set Input coordinates to the autopick.star file from step 15, set the additional parameters below, and click theRun! button.
Figure 8. Determining crossover distance, helical twist, and helical rise. A. An initial map depicts the crossover distance observed in twisting fibrils. The crossover distance is described as the length where the fibril turns 180° (red dotted line). Scale bar, 100 nm. B. The crossover distance can be measured (red line) from well-aligned 2D classes where the twisting nature of the fibril is observed; this requires a box size that spans a distance that is close to or larger than the crossover distance for an accurate measurement to be made. Here, a box size of 864 pixels (720 Å) was used for initial crossover estimates. Poor 2D classes that are misaligned or blurry prevent crossover distance measurements. C. The helical rise can be determined from 2D classes with a small box size (360 pixels) extracted at their original pixel size (0.834 Å/pix) that yield high-resolution details (i.e., spacing of the β-sheets). The sigma contrast of the 2D classes must be adjusted to visualize the helical layer lines in reciprocal space. From the average power spectrum, a measurement (red line) can be made from the meridian to the highest intensity layer line. This measurement can be used to estimate the helical rise. D. The measurements made in B and C are used to calculate the helical rise and the crossover distance. Then, the crossover distance and helical rise are used to calculate the helical twist of the structure. The estimated helical parameters are used for subsequent 3D refinement steps.
I/O:
Micrograph STAR file: Select/job004/micrographs.star
Input coordinates: AutoPick/job041/autopick.star
OR re-extract refined particles? No
OR re-center refined coordinates? No
Write output in float16? Yes
Extract:
Particle box size (pix): 864
Invert contrast? Yes
Normalize particles? Yes
Diameter background circle (pix): -1
Stddev for white dust removal: -1
Stddev for black dust removal: -1
Rescale particles? Yes
Re-scale size (pixels): 144
Use autopick FOM threshold? No
Helix:
Extract helical segments? Yes
Tube diameter (Å): 140
Use bimodal angular priors? Yes
Coordinates are start-end only? Yes
Cut helical tubes into segments? Yes
Number of unique asymmetrical units: 15
Helical rise (Å): 4.82
This job results in 771,754 particles with an original box size of 864 pixels that is rescaled to 144 pixels at a pixel size of 5.004 Å/pixel.
Note: The number of asymmetrical units was increased to 15. This results in an interbox distance of ~72 Å or ~25% of the small box size (360 pixels) that will be used for the final reconstruction.
17. 2D classification (large box size)
Classify the particles to remove junk particles and to estimate the crossover distance. Select the 2D classification job, set Input images STAR file to the particles.star file from step 16, set the additional parameters below, and then click on the Run! button.
I/O:
Input images STAR file: Extract/job042/particles.star
CTF:
Do CTF-correction? Yes
Ignore CTFs until first peak? Yes
Optimisation:
Number of classes: 50
Regularisation parameter T: 2
Use EM algorithm? Yes
Number of EM iterations: 20
Use VDAM algorithm? No
Mask diameter (Å): 710
Mask individual particles with zeros? Yes
Limit resolution E-step to (Å): -1
Center class averages? Yes
Sampling:
Perform image alignment? Yes
In-plane angular sampling: 2
Offset search range (pix): 5
Offset search step (pix): 1
Allow coarser sampling? No
Helix:
Classify 2D helical segments? Yes
Tube diameter (Å): 140
Do bimodal angular searches? Yes
Angular search range-psi (deg): 6
Restrict helical offsets to rise: Yes
Helical rise (Å): 4.82
This job results in therun_it020_optimiser.star file that contains the 2D class averages. This file can be viewed using theDisplay: drop-down menu on the right side of the GUI.
Sometimes, it can be helpful to determine the helical rise of the filament rather than assume 4.8 Å as the starting point. To do this, users can utilize 2D classifications and measurements of the average power spectra in Fourier space to calculate the estimated rise. To perform this analysis, use a box size of 360 pixels and high-resolution data (0.834 Å/pix), as this allows for more detail to be visualized in the 2D classes (specifically the β-sheet rungs). To do so, use the Particle Extraction job to extract particles to their original pixel size. Use the parameters as instructed in step 16, but ensure that Particle box size is set to 360 and that Rescale particles is set toNo. Once particle extraction is complete, run a 2D Classification job as described in step 17. Ensure Input images STAR files is set to the correct particles.star file from the Particle Extraction job and Mask diameter is set to 300. When the job is done, open the average power spectra by selecting the out:run_it020_optimiser from the display output. Enter an increased Sigma Contrast value in the top box of the RELION display GIU (we used 1 for our data) (Figure 8C). If the user fails to increase the Sigma Contrast, the average power spectra will not be visible (Figure 8C). Once the 2D classes are displayed, right-click on a class and select Show Fourier amplitudes (2X). This will open an image of the average power spectra. Make a measurement from the meridian to either layer line with the strong intensity (Figure 8C). This can be done by clicking and holding the center button on the mouse. Use the following formula to calculate the rise:
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(Figure 8C, 8D). For new experimental data, if the rise is substantially different, then parameters for steps 16 onward should reflect the updated rise. Here, a measurement of ~124 pixels results in a helical rise of 4.84 Å that will be refined in later steps (Figure 8C).
18. Subset selection (2D classes for initial map)
Select 2–3 good classes that will be used to generate an initial 3D volume. Select the Subset Select job, set Select classes from job to the run_it020_optimiser.star file from step 17, set the parameters below, then click the Run! button. A RELION display GUI will appear; reverse sort the class averages by rlnClassDistribution (as described in step 11) and select 2 class averages (Figure 9A, green boxes). To measure the crossover distance, right-click on a 2D class and select Show original image. A new window will appear. Using the center button, click and drag to measure the distance between two crossovers (Figure 8B). The distance in pixels is displayed over the image and in the terminal (Figure 8B). Multiply the measured distance by the current pixel size of 5.004 Å/pixel to calculate the distance in angstroms (Figure 8D). Here, we estimated a crossover distance of 120 pixels or 600 Å (Figure 8B). When done, close the original image. Repeat the process for any additional 2D classes you want to measure. Lastly, in the window with all the 2D classes, right-click and select Save STAR with selected images, and then close the display window.
Figure 9. Initial model generation. A. Classes selected from all 2D classes. All classes shown in A are the classes selected (job 21) from the classes rendered from the trained neural network auto-picking job on all micrographs (step 17). The green boxes indicate the two classes selected for initial model generation (step 18). B–G. Initial maps for the crossover distances 550–800 Å. One shows the cross-section of the refined filament (cross-section location shown by the black crossbar) and the other depicts the entirety of the filament. The commands used to generate (step 19) and rescale (step 20) the initial models are shown.
I/O:
Select classes from job: Class2D/job044/run_it020_optimiser.star
OR select from micrograph.star: Leave blank
OR select from particles.star: Leave blank
Class options:
Automatically select 2D classes? No
Re-center the class averages? Yes
Regroup the particles? No
Subsets:
Select based on metadata values? No
OR: select on image statistics? No
OR: split into subsets? No
Duplicates:
OR: remove duplicates? No
This job results in aclass_averages.star file containing the two selected classes.
19. Initial map generation usingrelion_helix_inimodel2d
Generate an initial map from the selected 2D classes in step 18 using the relion_helix_inimodel2d program [45]. The following steps must be completed in the terminal. First, create two directories to keep our data organized. In the terminal, navigate to the RELION project directory (this is the directory that contains all the RELION subdirectories) and enter the commands mkdir inimodel andmkdir ini4refine. This will create two directories: one that will house the initial volumes and a second that will contain the rescaled volumes that will be used for refinement steps. Documentation on relion_helix_inimodel2d can be found at https://relion.readthedocs.io/en/release-4.0/Reference/Helix.html. For convenience, we have detailed each argument below; alternatively, running relion_helix_inimodel2d with no additional arguments will detail all available arguments for the program. Before running the command, ensure that the input STAR file (--i) and the output root name (--o) are updated to your specific project and then run the command from the terminal. The following command generates an initial volume with an estimated crossover distance of 750 Å (Figure 9F).
relion_helix_inimodel2d --i Select/job045/class_averages.star --angpix 5.004 --mask_diameter 300 --sym 2 --iter 10 --search_shift 70 --search_angle 15 --search_size 10 --j 20 --crossover_distance 750 --o inimodel/Select045_CO750
The arguments run with relion_helix_inimodel2d are detailed below.
--i input STAR file with 2D classes
--angpix pixel size in angstroms
--mask_diameter size in angstroms of circular mask around 2D classes
--sym order of symmetry in 2D slices
--iter number of iterations to run
--search_shift distance in angstroms to search translations perpendicular to helical axis
--search_angle degrees to search in-plane rotations
--search_size ± number of pixels to fix best crossover distance
--j number of threads
--crossover_distance distance in angstroms between 2 crossovers
--o output root name
The program generates several files, and the initial 3D volume is saved with the suffix_class001_rec3d.mrc, which can be opened in ChimeraX for visualization [41,42,46]. Since the initial crossover distance is an estimate, we prefer to generate several initial maps for a round of 3D refinement to see what best fits our experimental data (Figure 9B–9G). Although our initial estimate for crossover distance was 600 Å, we found that an initial map with a crossover distance of 750 Å is best for this dataset (Figure 9C, 9F). You may test additional crossover distances as we typically do with new experimental datasets. To do so, change the --crossover_distance and --o arguments of the command above to generate additional maps of varying crossover distances with appropriate output root names (Figure 9B–9G).
20. Rescale initial map using relion_image_handler
The initial maps generated in the previous step must be rescaled because the 3D refinement steps will be performed with a smaller box size (360 pixels) at the original pixel size (0.834 Å/pixel). Use relion_image_handler to rescale the maps. For a list of all possible arguments, simply run the program in the terminal with no additional arguments. The command below was used to rescale the 750 Å crossover map. Before running the command, ensure the MRC input file (--i) and the MRC output file (--o) reflect your project. For convenience, the arguments used to runrelion_image_handler are detailed below.
relion_image_handler --i inimodel/Select045_CO750_class001_rec3d.mrc --angpix 5.004 --rescale_angpix 0.834 --new_box 360 --o ini4refine/Select045_CO750_box360.mrc
--i input MRC file of the initial map
--angpix pixel size in angstroms of the input file
--rescale_angpix scale input map to this new pixel size in angstroms
--new_box resize the input map to this box size in pixels
--o output name of resized map
Repeat this step for any additional maps that will be tested. Ensure that the input file (--i) and output file (--o) are updated to reflect the maps being rescaled (Figure 9B–9G).
21. Subset selection (2D classes for refinement)
Select additional classes from the 2D Classification job (step 17) to ensure there are enough particles for additional processing. Repeat the Subset Selection job as in step 18 but now select all the good classes for further processing (Figure 9A).
I/O:
Select classes from job: Class2D/job044/run_it020_optimiser.star
OR select from micrograph.star: Leave blank
OR select from particles.star: Leave blank
Class options:
Automatically select 2D classes? No
Re-center the class averages? Yes
Regroup the particles? No
Subsets:
Select based on metadata values? No
OR: select on image statistics? No
OR: split into subsets? No
Duplicates:
OR: remove duplicates? No
Here, we selected 21 classes with 413,249 particles saved in theparticles.star file (Figure 9A).
22. Particle extraction (small box size)
Select the Particle Extraction job, set Refined particles STAR files to the particles.star file from step 21, set the additional parameters below, and click the Run! button.
I/O:
Micrograph STAR file: Select/job004/micrographs.star
Input coordinates: Leave blank
OR re-extract refined particles? Yes
Refined particles STAR file: Select/job046/particles.star
Reset the refined offsets to zero? Yes
OR re-center refined coordinates? No
Write output in float16? Yes
Extract:
Particle box size (pix): 360
Invert contrast? Yes
Normalize particles? Yes
Diameter background circle (pix): -1
Stddev for white dust removal: -1
Stddev for black dust removal: -1
Rescale particles? No
Use autopick FOM threshold? No
Helix:
Extract helical segments? Yes
Tube diameter (Å): 140
Use bimodal angular priors? Yes
Coordinates are start-end only? Yes
Cut helical tubes into segments? Yes
Number of unique asymmetrical units: 15
Helical rise (Å): 4.82
The 413,249 particles were re-extracted to a box size of 360 pixels and a pixel size of 0.834 Å/pixel. The particles are stored in the particles.star file.
23. 3D auto-refine (fixed symmetry)
The particle set from step 22 and the rescaled initial map generated in step 20 will be subjected to a round of 3D refinement. First, take the estimated helical rise and calculate the initial twist for the estimated crossover distance using the following formula:
t
w
i
s
t
=
r
i
s
e
×
180
c
r
o
s
s
o
v
e
r
d
i
s
tan
c
e
(Figure 8D). The rise is estimated to be 4.82 Å, and the crossover distance was estimated to be 750 Å, so the initial twist value is 1.16°. Finally, apply a negative value to the initial twist based on the assumption that fibrils typically display a left-handed helical form, as supported by atomic force microscopy studies [13]. Select the 3D Auto-Refine job, set Input images STAR file to the particles.star file generated in step 22, set Reference map to the rescaled initial volume generated in step 20 (in our case, this file was named Select045_CO750_box360.mrc), set the parameters below, and click the Run! button.
I/O:
Input images STAR file: Extract/job047/particles.star
Reference map: ini4refine/Select045_CO750_box360.mrc
Reference mask (optional): Leave blank
Reference:
Ref. map is on absolute greyscale? No
Initial low-pass filter (Å): 10
Symmetry: C1
CTF:
Do CTF-correction? Yes
Ignore CTFs until first peak? No
Optimisation:
Mask diameter (Å): 220
Mask individual particles with zeros? Yes
Use solvent-flattened FSCs? No
Auto-sampling:
Initial angular sampling: 3.7 degrees
Initial offset range (pix): 5
Initial offset step (pix): 1
Local searches from auto-sampling: 1.8 degrees
Relax symmetry: Leave blank
Use finer angular sampling faster? (No)
Helix:
Do helical reconstruction? Yes
Tube diameter – inner, outer (Å): -1, 140
Angular search range – rot, tilt, psi (deg): -1, 15, 10
Range factor of local averaging: -1
Keep tilt-prior fixed: Yes
Apply helical symmetry? Yes
Number of unique asymmetrical units: 15
Initial twist (deg), rise (Å): -1.16, 4.82
Central Z length (%): 25
Do local searches of symmetry? No
The job results in a map with a global resolution of 3.66 Å (Figure 10A, 10B, blue). Repeat this step for any additional initial maps and crossover distances that you would like to test. We tested crossover distances of 550, 600, 650, 700, 750, and 800 Å (Figure 10A, 10B). These tests showed that three maps resolved to a resolution of 3.66 Å. The map generated from the 750 Å crossover distance was selected because the backbone density was best resolved, and the map showed side chain densities for some residues (Figure 10B, blue). Additionally, the map showed a clear separation of the b-strands along the helical axis.
Figure 10. 3D refinement of different crossover distances and 3D classification. A. Cross-sections, resolution, and calculated twist and rise of each initial model after 3D refinement (550–800 Å) (step 23). Red and blue squares indicate respective electron potential maps for B. B. Cross-section of the electron potential maps refined with 600 Å (red) and 750 Å (blue) crossovers. Scale bar, 25 Å. C. 3D classifications from 750 Å crossover initial model (step 24). The green box indicates the selected 3D class used for further refinement (step 25).
24. 3D classification (symmetry search)
During the Subset Selection job (step 21), we selected all the 2D classes that resembled amyloid fibrils. Being less stringent after 2D classification means that heterogeneity most likely exists in our dataset. By using 3D classification, we can further sort the heterogeneity that may exist in the particle set and improve the quality of the reconstruction. Use the 3D reconstruction from step 23 as an initial starting point to then sort particles into four classes. Use the Do local searches of symmetry tool to search a range of helical parameters that best fit the dataset. Select the 3D Classification job, set Input images STAR file to the run_data.star file from step 23, set Reference map to the run_half1_class001_unfil.mrc from step 23, set the additional parameters below, and click theRun! button.
I/O:
Input images STAR file: Refine3D/job069/run_data.star
Reference map: Refine3D/job069/run_half1_class001_unfil.mrc
Reference mask (optional): Leave blank
Reference:
Ref. map is on absolute greyscale? No
Initial low-pass filter (Å): 4.5
Symmetry: C1
CTF:
Do CTF-correction? Yes
Ignore CTFs until first peak? No
Optimization:
Number of classes: 4
Regularization parameter T: 4
Number of iterations: 20
Use fast subsets (for large data sets)? No
Mask diameter (Å): 220
Mask individual particles with zeros? Yes
Limit resolution E-step to (Å): -1
Sampling:
Perform image alignment? Yes
Angular sampling interval: 3.7 degrees
Offset search range (pix): 5
Offset search step (pix): 1
Perform local angular searches? No
Allow coarser sampling? No
Helix:
Do helical reconstruction? Yes
Tube diameter – inner, outer (Å): -1, 140
Angular search range – rot, tilt, psi (deg): -1, 15, 10
Range factor of local averaging: -1
Keep tilt-prior fixed: Yes
Apply helical symmetry? Yes
Number of unique asymmetrical units: 15
Initial twist (deg), rise (Å): -1.14, 4.82
Central Z length (%): 25
Do local searches of symmetry? Yes
Twist search – Min, Max, Step (deg): -0.9, -1.2, 0.01
Rise search – Min, Max, Step (Å): 4.75, 4.95, 0.01
The job runs for 20 iterations, sorting the particle set into four classes and optimizing helical parameters at each iteration. A cross-section of the 3D volumes can be visualized by displaying the run_it020_optimiser.star file in RELION. Alternatively, the four MRC files generated in this job (run_it020_class001.mrc, run_it020_class002.mrc, etc.) can be opened in ChimeraX for easier visualization of the 3D maps. Class 3 was the best 3D volume with a helical twist of -1.12° and a helical rise of 4.84 Å (Figure 10C, green box).
25. Subset selection (3D class for additional processing)
Use the Subset selection job to select the best class from the 3D classification job in step 24. Ensure Select classes from job is set to the run_it020_optimiser.star file that was generated in step 24. Set the parameters below and click the Run! button. Refer to step 12 for how to display, select, and save classes in a Subset selection job.
I/O:
Select classes from job: Class3D/job077/run_it020_optimiser.star
OR select from micrograph.star: Leave blank
OR select from particles.star: Leave blank
Class options:
Automatically select 2D classes? No
Re-center the class averages? Yes
Regroup the particles? No
Subsets:
Select based on metadata values? No
OR: select on image statistics? No
OR: split into subsets? No
Duplicates:
OR: remove duplicates? No
Class 3 was selected in this job, and the data was saved to the particles.star file that contained all 129,895 particles for that class (Figure 10C).
26. 3D auto-refine (symmetry search)
Select the 3D auto-refine job and update Input images STAR file to the particles.star file from step 25 and the Reference map to the best 3D map from the 3D classification in step 24 (in our case, this was run_it020_class003.mrc, but it may be different for your project). Then, set the parameters below and click the Run! button.
I/O:
Input images STAR file: Select/job080/particles.star
Reference map: Class3D/job077/run_it020_class003.mrc
Reference mask (optional): Leave blank
Reference:
Ref. map is on absolute greyscale? No
Initial low-pass filter (Å): 4.5
Symmetry: C1
CTF:
Do CTF-correction? Yes
Ignore CTFs until first peak? No
Optimisation:
Mask diameter (Å): 220
Mask individual particles with zeros? Yes
Use solvent-flattened FSCs? No
Auto-sampling:
Initial angular sampling: 3.7 degrees
Initial offset range (pix): 5
Initial offset step (pix): 1
Local searches from auto-sampling: 1.8 degrees
Relax symmetry: Leave blank
Use finer angular sampling faster? (No)
Helix:
Do helical reconstruction? Yes
Tube diameter – inner, outer (Å): -1, 140
Angular search range – rot, tilt, psi (deg): -1, 15, 10
Range factor of local averaging: -1
Keep tilt-prior fixed: Yes
Apply helical symmetry? Yes
Number of unique asymmetrical units: 15
Initial twist (deg), rise (Å): -1.11, 4.84
Central Z length (%): 25
Do local searches of symmetry? Yes
Twist search – Min, Max, Step (deg): -0.9, -1.3, 0.01
Rise search – Min, Max, Step (Å): 4.75, 4.95, 0.01
The optimized helical parameters converged to a helical twist of -1.11° and a rise of 4.84 Å. The resolution without masking is 3.23 Å. Therun_class001.mrc file can be downloaded and opened with ChimeraX to visualize the 3D volume (Figure 11, step 26).
Figure 11. Results of 3D refinements and post-processing steps. The 3D refinements and their corresponding post-processed maps of our processing pipeline are depicted here. The step number, resolution, twist, rise, and mask percentages are displayed for each electron potential map. A description as to whether the electron potential map display is a result of a 3D refinement job or a post-processing job is displayed at the top of the figure. The processing workflow incrementally improves maps’ quality and resolution, resulting in a final map at 2.04 Å resolution.
27. Mask creation (80% mask)
Helical reconstruction is prone to loss of resolvability as the volume reaches the edge of the box. Thus, masking encompasses a central portion of the fibril and excludes the ends of the fibril. The mask can be as small as the Central Z length established in the 3D auto-refine job. However, at this stage in processing, we may benefit from a larger mask to ensure we have sufficient signal for the CTF refinement steps. Open the run_class001.mrc file from step 26 in ChimeraX. Ensure that the volume step is set to 1; then, lower the volume threshold until noise starts to appear in the solvent space. Note this threshold and set this as the Initial binarization threshold for the Mask creation job. A value of 0.00096 worked well for this project. Update the Input 3D map to the run_class001.mrc generated in step 26. Set the additional parameters below and then click the Run! button.
I/O:
Input 3D map: Refine3D/job081/run_class001.mrc
Mask:
Lowpass filter map (Å) 15
Pixel size (Å) -1
Initial binarization threshold: 0.00096
Extend binary map this many pixels: 5
Add a soft-edge of this many pixels: 5
Helix:
Mask a 3D helix? Yes
Central Z length (%): 80
In ChimeraX, open the mask.mrc file and the run_class001.mrc file from step 26. Ensure both maps are set to a step size of 1, set the mask threshold to 0.99 to visualize the mask volume, and, for easier visualization, lower the mask opacity to 50% (Figure 12A). Inspect the mask and map; when viewing the central cross-section of the map, ensure the entire proteinaceous volume is within the mask. If there are no issues, then proceed to the next step. However, if the map is not completely encompassed by the mask, lower the Initial binarization threshold value and rerun the job by clicking the Continue! button. Repeat this process until the mask is satisfactory (Figure 12A).
Figure 12. Mask central Z length coverage. A. A mask (gray) covering 80% of the map (purple) along the fibril axis (step 27) used during CTF refinement steps. B. A mask (gray) covering 25% of the map (purple) along the fibril axis (step 44) used in the final post-processing job (step 45). C. Filament after applying real-space symmetrization (step 47) to the edge of the box using the relion_helix_toolbox program. Scale bars, 25 Å.
28. Post-processing
The post-processing job will recalculate the global resolution with masking and will automatically estimate and apply a B-factor to sharpen the map, further improving the quality of the map. Select the Post-processing job, set One of the 2 unfiltered half-maps to the run_half1_class001_unfil.mrc file from step 26, set Solvent mask to the mask.mrc file from step 27, and set MTF of the detector (STAR file) to the k3-CDS-300keV-mtf.star file that is supplied with EMPIAR-12229. Set the remaining parameters below and then click the Run! button.
I/O:
One of the 2 unfiltered half-maps: Refine3D/job081/run_half1_class001_unfil.mrc
Solvent mask: MaskCreate/job086/mask.mrc
Calibrated pixel size (Å) -1
Sharpen:
Estimate B-factor automatically? Yes
Lowest resolution for auto-B fit (Å): 10
Use your own B-factor? No
Skip FSC-weighting? No
MTF of the detector (STAR file): k3-CDS-300keV-mtf.star
Original detector pixel size: -1
The job estimated a B-factor of -97, and the processed map is saved as postprocess.mrc. The job also calculated a resolution of 2.97 Å with masking, and the volume is saved as postprocess_mask.mrc (Figure 11, step 28).
29. Bayesian polishing (round 1)
The next steps will aim at improving the quality of the particles to further improve the resolvability of the map. The polishing will use motion-corrected micrographs and particle positions to improve motion correction on a per-particle basis. Select the Bayesian polishing job, set the Micrographs (from MotionCorr) to the corrected_micrographs.star file from step 2, set the Particles (from Refine 3D or CtfRefine) to the run_data.star file from step 26, set the Postprocess STAR file to thepostprocess.star file from step 28, set the remaining parameters below, and click the Run! button.
I/O:
Micrographs (from MotionCorr): MotionCorr/job002/corrected_micrographs.star
Particles (from Refine 3D or CtfRefine): Refine3D/job081/run_data.star
Postprocess STAR file: PostProcess/job088/postprocess.star
First movie frame: 1
Last movie frame: -1
Extraction size (pix in unbinned movie): -1
Re-scale size (pixels): -1
Write output in float16? Yes
Train:
Train optimal parameters? No
Polish:
Perform particle polishing? Yes
Optimized parameter file: Leave blank
OR use your own parameters?
Sigma for velocity (Å/dose): 0.2
Sigma for divergence (Å): 5000
Sigma for acceleration (Å/dose): 2
Minimum resolution for B-factor fit (Å): 20
Maximum resolution for B-factor fit (Å): -1
The job will save the particles to the shiny.star file. The improvements of the particle positions can be found in the logfile.pdf file.
30. 3D auto-refine (pseudo-screw symmetry)
Up to this point, we have only applied helical symmetry to the 3D reconstruction. We will now address additional symmetry that may be present to further improve the quality of the reconstruction. Previous studies have shown that amyloid fibrils exist with varying degrees of symmetry. For two protofilament fibrils, we observe either C2 symmetry, where two protofilaments are identical and in register, as commonly observed in Tau fibrils, or we observe pseudo-screw symmetry (P21), where two protofilaments are identical but out of register, as observed in a-synuclein fibrils (Figure 1G, 1F) [12,13,47]. To understand this difference in symmetry, it is necessary to manually inspect the reconstruction to determine the best symmetry for the dataset. This can be done by using ChimeraX to analyze the 3D volume from either the run_class001.mrc file from step 26 or the postprocess.mrc file from step 28. Here, we determined that pseudo-screw symmetry exists within our dataset. To apply this symmetry, we will continue to set the Symmetry parameter to C1, but we will divide the helical rise in half and subtract the helical twist from 180°. By doing so, we can impose pseudo-screw symmetry on our reconstruction. Select the 3D auto-refine job, set Input images STAR files to the shiny.star file generated in step 29, set Reference map to the run_half1_class001_unfil.mrc file from job 26, set the additional parameters below, and then click the Run! button.
I/O:
Input images STAR file: Polish/job091/shiny.star
Reference map: Refine3D/job081/run_half1_class001_unfil.mrc
Reference mask (optional): Leave blank
Reference:
Ref. map is on absolute greyscale? No
Initial low-pass filter (Å): 4.5
Symmetry: C1
CTF:
Do CTF-correction? Yes
Ignore CTFs until first peak? No
Optimization:
Mask diameter (Å): 220
Mask individual particles with zeros? Yes
Use solvent-flattened FSCs? No
Auto-sampling:
Initial angular sampling: 3.7 degrees
Initial offset range (pix): 5
Initial offset step (pix): 1
Local searches from auto-sampling: 1.8 degrees
Relax symmetry: Leave blank
Use finer angular sampling faster? (No)
Helix:
Do helical reconstruction? Yes
Tube diameter – inner, outer (Å): -1, 140
Angular search range – rot, tilt, psi (deg): -1, 15, 10
Range factor of local averaging: -1
Keep tilt-prior fixed: Yes
Apply helical symmetry? Yes
Number of unique asymmetrical units: 15
Initial twist (deg), rise (Å): 179.445, 2.42
Central Z length (%): 25
Do local searches of symmetry? Yes
Twist search – Min, Max, Step (deg): 179.24, 179.65, 0.01
Rise search – Min, Max, Step (Å): 2.2, 2.6, 0.01
The unmasked reconstruction improved from 3.23 Å (step 26) to 3.00 Å and is stored in therun_class001.mrc file (Figure 11, step 30). The symmetry parameters reflect pseudo-screw symmetry and were optimized to a twist of 179.45° and a rise of 2.42 Å.
31. Post-processing
Run a Post-processing job to see how masking the solvent region improves the resolution and how automated sharpening can improve the map quality. Select the Post-processing job, set One of the 2 unfiltered half-maps to the run_half1_class001_unfil.mrc file from step 30, set the Solvent mask to the mask.mrc file from step 27, set the additional parameters below, and then click the Run! button.
I/O:
One of the 2 unfiltered half-maps: Refine3D/job093/run_half1_class001_unfil.mrc
Solvent mask: MaskCreate/job086/mask.mrc
Calibrated pixel size (Å) -1
Sharpen:
Estimate B-factor automatically? Yes
Lowest resolution for auto-B fit (Å): 10
Use your own B-factor? No
Skip FSC-weighting? No
MTF of the detector (STAR file): k3-CDS-300keV-mtf.star
Original detector pixel size: -1
Use ChimeraX to visualize the improvements to the postprocess_masked.mrc map. The GS-FSC0.143 for this map with masking improved from 2.97 to 2.89 Å, and sharpening improved side-chain densities throughout the map (Figure 11, step 31).
32. CTF refinement (anisotropic magnification, round 1)
The next three steps will utilize the CTF refinement job to improve the CTF fits for the particle set. The three jobs perform corrections, namely 1) anisotropic magnification, 2) asymmetrical and symmetrical aberrations, and 3) recalculation of per-particle defocus and per-micrograph astigmatism. Together, these steps improve CTF fits that translate into improvements in the reconstruction. The first job will correct for anisotropic magnification. Select the CTF refinement job, set the Particles (from Refine3D) to the run_data.star file from step 30, set the Postprocess STAR file to the postprocess.star file from step 31, set the parameters below, and then click the Run! button.
I/O:
Particles (from Refine3D): Refine3D/job093/run_data.star
Postprocess STAR file: PostProcess/job095/postprocess.star
Fit:
Estimate (anisotropic) magnification? Yes
Minimum resolution for fits (Å): 30
This job estimated a magnification anisotropy of 0.31% and stored the refined particles in the particles_ctf_refine.star file.
33. CTF refinement (asymmetrical and symmetrical aberrations, round 1)
Use the refined particles from the previous job to correct for asymmetrical and symmetrical aberrations. Select the CTF refinement job, set Particles (from Refine3D) to the particles_ctf_refine.star file from step 32, set Postprocess STAR file to the postprocess.star file from step 31, set the parameters below, and then click the Run! button.
I/O:
Particles (from Refine3D): CtfRefine/job096/particles_ctf_refine.star
Postprocess STAR file: PostProcess/job095/postprocess.star
Fit:
Estimate (anisotropic) magnification? No
Perform CTF parameter fitting? Yes
Fit defocus? Per-particle
Fit astigmatism? Per-micrograph
Fit B-factor? No
Fit phase-shift? No
Estimate beamtilt? No
Estimate 4th order aberrations? No
Minimum resolution for fits (Å): 30
The refined particles are stored in the particles_ctf_refine.star file and the results of the job can be visualized by opening the logfile.pdf.
34. CTF refinement (recalculate defocus and astigmatism, round 1)
Next, recalculate defocus values on a per-particle basis and astigmatism on a per-micrograph basis. Select the CTF refinement job, set Particles (from Refine3D) to the particles_ct_refine.star file from step 33, set Postprocess STAR file to the postprocess.star file from step 31, set the additional parameters below, and then click the Run! button.
I/O:
Particles (from Refine3D): CtfRefine/job097/particles_ctf_refine.star
Postprocess STAR file: PostProcess/job095/postprocess.star
Fit:
Estimate (anisotropic) magnification? No
Perform CTF parameter fitting? No
Estimate beamtilt? Yes
Also estimate trefoil? Yes
Estimate 4th order aberrations? Yes
Minimum resolution for fits (Å): 30
The refined particles are saved to the particles_ctf_refine.star file and are now ready for 3D refinement.
35. 3D auto-refine (CTF refined particles, round 1)
Generate a new 3D volume with the refined particles. Select the 3D auto-refine job, set Input images STAR file to the particles_ctf_refine.star file from step 34, set Reference map to the run_half1_class001.mrc file from step 30, set the parameters below, ensure that the helical parameters are updated to the optimized twist and rise values from step 30 (these are found in the output log from step 30), and then click the Run! button.
I/O:
Input images STAR file: CtfRefine/job098/particles_ctf_refine.star
Reference map: Refine3D/job093/run_half1_class001_unfil.mrc
Reference mask (optional): Leave blank
Reference:
Ref. map is on absolute greyscale? No
Initial low-pass filter (Å): 4.5
Symmetry: C1
CTF:
Do CTF-correction? Yes
Ignore CTFs until first peak? No
Optimization:
Mask diameter (Å): 220
Mask individual particles with zeros? Yes
Use solvent-flattened FSCs? No
Auto-sampling:
Initial angular sampling: 3.7 degrees
Initial offset range (pix): 5
Initial offset step (pix): 1
Local searches from auto-sampling: 1.8 degrees
Relax symmetry: Leave blank
Use finer angular sampling faster? (No)
Helix:
Do helical reconstruction? Yes
Tube diameter – inner, outer (Å): -1, 140
Angular search range – rot, tilt, psi (deg): -1, 15, 10
Range factor of local averaging: -1
Keep tilt-prior fixed: Yes
Apply helical symmetry? Yes
Number of unique asymmetrical units: 15
Initial twist (deg), rise (Å): 179.448, 2.42
Central Z length (%): 25
Do local searches of symmetry? Yes
Twist search – Min, Max, Step (deg): 179.24, 179.65, 0.01
Rise search – Min, Max, Step (Å): 2.2, 2.6, 0.01
After CTF refinement, the resolution of the unmasked 3D reconstruction improved from 3.00 to 2.38 Å (Figure 11, step 35). The helical parameters did not change and converged to a twist of 179.45° and a rise of 2.42 Å. The 3D map is saved to the run_class001.mrc file and can be opened in ChimeraX for visualization.
36. Post-processing
Apply the mask from step 27 to recalculate the FSC and sharpen the map. Select the Post-processing job, set One of the 2 unfiltered half-maps to the run_half1_class001_unfil.mrc file from step 35, set Solvent mask to the mask.mrc file from step 27, set the additional parameters below, and click the Run! button.
I/O:
One of the 2 unfiltered half-maps: Refine3D/job099/run_half1_class001_unfil.mrc
Solvent mask: MaskCreate/job086/mask.mrc
Calibrated pixel size (Å) -1
Sharpen:
Estimate B-factor automatically? Yes
Lowest resolution for auto-B fit (Å): 10
Use your own B-factor? No
Skip FSC-weighting? No
MTF of the detector (STAR file): k3-CDS-300keV-mtf.star
Original detector pixel size: -1
The B-factor was estimated to -55 and applied to the map. The resolution of the masked map improved from 2.89 to 2.31 Å (Figure 11, step 36). The 3D map was saved to the postprocess_masked.mrc file and can be visualized in ChimeraX.
37. Bayesian polishing (round 2)
Perform one more cycle of polishing and CTF refinement before a final round of 3D refinement and postprocessing (steps 29–36). Select the Bayesian polishing job, set Micrographs (from MotionCorr) to the corrected_micorgraphs.star file from step 2, set Particles from Refine 3D or CtfRefine to the run_data.star file from step 35, set Postprocess STAR file to the postprocess.star file from step 36, set the additional parameters below, and click the Run! button.
I/O:
Micrographs (from MotionCorr): MotionCorr/job002/corrected_micrographs.star
Particles from Refine 3D or CtfRefine: Refine3D/job099/run_data.star
Postprocess STAR file: PostProcess/job100/postprocess.star
First movie frame: 1
Last movie frame: -1
Extraction size (pix in unbinned movie): -1
Re-scale size (pixels): -1
Write output in float16? Yes
Train:
Train optimal parameters? No
Polish:
Perform particle polishing? Yes
Optimized parameter file: Leave blank
OR use your own parameters?
Sigma for velocity (Å/dose): 0.2
Sigma for divergence (Å): 5000
Sigma for acceleration (Å/dose): 2
Minimum resolution for B-factor fit (Å): 20
Maximum resolution for B-factor fit (Å): -1
The polished particles are stored in theshiny.star file.
38. 3D auto-refine (polished particles, round 2)
Use the polished particles from step 37 and perform a round of 3D refinement. Select the 3D auto-refine job, set Input images STAR files to the shiny.star file from step 37, set Reference map to the run_half1_class001_unfil.mrc file from step 35, set the parameters below, and click the Run! button.
I/O:
Input images STAR file: Polish/job101/shiny.star
Reference map: Refine3D/job099/run_half1_class001_unfil.mrc
Reference mask (optional): Leave blank
Reference:
Ref. map is on absolute greyscale? No
Initial low-pass filter (Å): 4.5
Symmetry: C1
CTF:
Do CTF-correction? Yes
Ignore CTFs until first peak? No
Optimization:
Mask diameter (Å): 220
Mask individual particles with zeros? Yes
Use solvent-flattened FSCs? No
Auto-sampling:
Initial angular sampling: 3.7 degrees
Initial offset range (pix): 5
Initial offset step (pix): 1
Local searches from auto-sampling: 1.8 degrees
Relax symmetry: Leave blank
Use finer angular sampling faster? (No)
Helix:
Do helical reconstruction? Yes
Tube diameter – inner, outer (Å): -1, 140
Angular search range – rot, tilt, psi (deg): -1, 15, 10
Range factor of local averaging: -1
Keep tilt-prior fixed: Yes
Apply helical symmetry? Yes
Number of unique asymmetrical units: 15
Initial twist (deg), rise (Å): 179.449, 2.42
Central Z length (%): 25
Do local searches of symmetry? Yes
Twist search – Min, Max, Step (deg): 179.24, 179.65, 0.01
Rise search – Min, Max, Step (Å): 2.2, 2.6, 0.01
After a second round of polishing, the unmasked map did not improve in resolution, staying at 2.38 Å (Figure 11, step 38). Next, we will see if there is an improvement in the masked reconstruction.
39. Post-processing
Select the Post-processing job, set One of the 2 unfiltered half-maps to the run_half1_class001_unfil.mrc file from step 38, set the Solvent mask to the mask.mrc file from step 27, set the additional parameters below, and click the Run! button.
I/O:
One of the 2 unfiltered half-maps: Refine3D/job102/run_half1_class001_unfil.mrc
Solvent mask: MaskCreate/job086/mask.mrc
Calibrated pixel size (Å) -1
Sharpen:
Estimate B-factor automatically? Yes
Lowest resolution for auto-B fit (Å): 10
Use your own B-factor? No
Skip FSC-weighting? No
MTF of the detector (STAR file): k3-CDS-300keV-mtf.star
Original detector pixel size: -1
The resolution of the masked reconstruction increased slightly from 2.31 to 2.27 Å (Figure 11, step 39). The 3D map is stored in the postprocess_masked.mrc file and can be visualized in ChimeraX.
40. CTF refinement (anisotropic magnification, round 2)
Perform a final round of CTF refinements as in steps 32–34. Select the CTF refinement job, set Particles (from Refine3D) to the run_data.star file from step 38, set Postprocess STAR file to the postprocess.star file from step 39, set the parameters below, and then click the Run! button.
I/O:
Particles (from Refine3D): Refine3D/job102/run_data.star
Postprocess STAR file: PostProcess/job103/postprocess.star
Fit:
Estimate (anisotropic) magnification? Yes
Minimum resolution for fits (Å): 30
The refined particles are stored in the particles_ctf_refine.star file and will be used in the next step.
41. CTF refinement (asymmetrical and symmetrical aberrations, round 2)
Select the CTF refinement job, set Particles (from Refine3D) to the particles_ctf_refine.star file from step 40, set Postprocess STAR file to the postprocess.star file from step 39, set the parameters below, and then click the Run! button.
I/O:
Particles (from Refine3D): CtfRefine/job104/particles_ctf_refine.star
Postprocess STAR file: PostProcess/job103/postprocess.star
Fit:
Estimate (anisotropic) magnification? No
Perform CTF parameter fitting? Yes
Fit defocus? Per-particle
Fit astigmatism? Per-micrograph
Fit B-factor? No
Fit phase-shift? No
Estimate beamtilt? No
Estimate 4th order aberrations? No
Minimum resolution for fits (Å): 30
The particles were written out to theparticles_ctf_refine.star file and will be used in the next step.
42. CTF refinement (recalculate defocus and astigmatism, round 2)
Select the CTF refinement job, set Particles (from Refine3D) to the particles_ctf_refine.star file from step 41, set Postprocess STAR files to the postprocess.star file from step 39, set the additional parameters below, and then click the Run! button.
I/O:
Particles (from Refine3D): CtfRefine/job105/particles_ctf_refine.star
Postprocess STAR file: PostProcess/job103/postprocess.star
Fit:
Estimate (anisotropic) magnification? No
Perform CTF parameter fitting? No
Estimate beamtilt? Yes
Also estimate trefoil? Yes
Estimate 4th order aberrations? Yes
Minimum resolution for fits (Å): 30
The refined particles are stored in the particls_ctf_refine.star file and are ready for 3D refinement.
43. 3D auto-refine (CTF refined particles, round 2)
Run a 3D refinement using the CTF-refined particles. Select the 3D auto-refine job, set Input images STAR file to the particles_ctf_refine.star file from step 42, set Reference map to the run_half1_class001_unfil.mrc file from step 38, set the additional parameters below, and click the Run! button.
I/O:
Input images STAR file: CtfRefine/job106/particles_ctf_refine.star
Reference map: Refine3D/job102/run_half1_class001_unfil.mrc
Reference mask (optional): Leave blank
Reference:
Ref. map is on absolute greyscale? No
Initial low-pass filter (Å): 4.5
Symmetry: C1
CTF:
Do CTF-correction? Yes
Ignore CTFs until first peak? No
Optimization:
Mask diameter (Å): 220
Mask individual particles with zeros? Yes
Use solvent-flattened FSCs? No
Auto-sampling:
Initial angular sampling: 3.7 degrees
Initial offset range (pix): 5
Initial offset step (pix): 1
Local searches from auto-sampling: 1.8 degrees
Relax symmetry: Leave blank
Use finer angular sampling faster? (No)
Helix:
Do helical reconstruction? Yes
Tube diameter – inner, outer (Å): -1, 140
Angular search range – rot, tilt, psi (deg): -1, 15, 10
Range factor of local averaging: -1
Keep tilt-prior fixed: Yes
Apply helical symmetry? Yes
Number of unique asymmetrical units: 15
Initial twist (deg), rise (Å): 179.449, 2.42
Central Z length (%): 25
Do local searches of symmetry? Yes
Twist search – Min, Max, Step (deg): 179.24, 179.65, 0.01
Rise search – Min, Max, Step (Å): 2.2, 2.6, 0.01
The resolution of the unmasked map increased slightly from 2.38 to 2.35 Å (Figure 11, step 43). This result suggests that any additional rounds of polishing or CTF refinement will not yield meaningful gains in map quality and are thus not necessary.
44. Mask creation (25% mask)
In the Helix tab of the 3D auto-refine job, we set the central Z length to 25% of the particle box. This central region is where searching for helical symmetry occurs and is also the region where real-space helical symmetry is imposed. In the previous Mask creation job from step 27, the mask length was set to 80% of the central axis to ensure enough signal was available for CTF refinements. Now, in the final stages of processing, we can reduce the mask size to a central Z length of 25% as was used in the 3D reconstruction steps. As in step 27, you may need to open the run_class001.mrc file from step 43 in ChimeraX to determine the appropriate Initial binarization threshold for the reconstruction. A value of 0.0011 worked well for us. Select the Mask creation job, set Input 3D map to the run_class001.mrc file from step 43, set the parameters below, and then click the Run! button.
I/O:
Input 3D map: Refine3D/job107/run_class001.mrc
Mask:
Lowpass filter map (Å) 15
Pixel size (Å) -1
Initial binarization threshold: 0.0011
Extend binary map this many pixels: 5
Add a soft-edge of this many pixels: 5
Helix:
Mask a 3D helix? Yes
Central Z length (%): 25
The mask is saved to the mask.mrc file and will be used in the next step (Figure 12B).
45. Post-processing
Apply the latest mask from step 44 to the final reconstruction from step 43 to recalculate the resolution and B-factor. Select the Post-processing job, set One of the 2 unfiltered half-maps to the run_half1_class001_unfil.mrc file from step 43, set Solvent mask to the mask.mrc file from step 44, set the additional parameters below, and then click the Run! button.
I/O:
One of the 2 unfiltered half-maps: Refine3D/job107/run_half1_class001_unfil.mrc
Solvent mask: MaskCreate/job109/mask.mrc
Calibrated pixel size (Å) -1
Sharpen:
Estimate B-factor automatically? Yes
Lowest resolution for auto-B fit (Å): 10
Use your own B-factor? No
Skip FSC-weighting? No
MTF of the detector (STAR file): k3-CDS-300keV-mtf.star
Original detector pixel size: -1
The final masked map has a resolution of 2.04 Å and a B-factor of -43 (Figure 11, step 45). The map displays well-resolved side chain densities as expected for a map at ~2 Å resolution.
Note: We observe a spike in the FSC plot at ~2.4 Å (the repeating unit) in both our reconstruction and in several published structures (Figure 13) [22,27,48]. This spike is alleviated with masking, but it is a common feature observed in amyloid structures that resolve to high resolution. Additionally, other helical structures, such as tad pili, also display a similar spike due to the strong signal at the repeating unit of ~4.9 Å [49].
Figure 13. Comparison of Fourier shell correlation (FSC) plots of a-synuclein maps deposited to the EMDB resolving to below 2.3 Å. The unmasked FSC plots (calculated FSC from deposited half maps, orange) for the deposited maps display an FSC spike at a spatial frequency of 0.4 Å-1 (~2.4 Å). The masked FSC plots (author provided FSC, blue) dampen this feature.
46. Local resolution
Calculate a local resolution map to understand the differences in resolution across the map. Select the Local resolution job, set One of the 2 unfiltered half-maps to the run_half1_class001_unfil.mrc file from step 43, set User-provided solvent mask to the mask.mrc file from step 44, set the additional parameters below, and then click the Run! button.
I/O:
One of the 2 unfiltered half-maps: Refine3D/job107/run_half1_class001_unfil.mrc
User-provided solvent mask: MaskCreate/job109/mask.mrc
Calibrated pixel size (Å): 0.834
ResMap:
Use ResMap? No
Relion:
Use Relion? Yes
User-provided B-factor: -40
MTF of the detector (STAR file): k3-CDS-300keV-mtf.star
The job results in a histogram.pdf file that contains a graph of the local resolution within the provided mask. The relion_locres.mrc file can be opened in ChimeraX along with the postprocess.mrc file from step 45 to color the surface of the map by resolution (Figure 14). Please see the “Analyzing the results” section in the RELION local resolution documentation page for details on handling these maps in ChimeraX (https://relion.readthedocs.io/en/latest/SPA_tutorial/Validation.html).
Figure 14. Local resolution map of a-syn fibril from cryo-EM data. A. Local resolution map of filamentous a-syn depicting a loss of resolution toward the end of the fibril, with the best resolution located along the central portion of the map. B. Cross-section of the local resolution map of filamentous a-syn showing that the best-resolved regions of the map are located along the fibril core and protofilament interface. Map resolution key spans from 1.8 Å (cyan) to 3.0 Å (red).
47. Real-space symmetrization (optional)
As stated previously, real-space symmetry is applied to only the central 25% of the reconstruction, and the molecular model is built into this central region (Figure 1E). However, in some cases, it is beneficial to extend the symmetrization to the edge of the box. For example, to better visualize the crossover distance, we generate a map with real-space symmetry imposed to the edge of the box, and then we align several models in ChimeraX to generate a multi-map volume that spans close to 1000 Å (Figure 1A). This process allows for easier visualization of the crossover distance when making figures. To impose real-space symmetry, run the relion_helix_toolbox command in the terminal. Before running the command, cd to the job directory for step 45.
relion_helix_toolbox --impose --i postprocess_masked.mrc --o postprocess_masked_sym.mrc --cyl_outer_diameter 220 --angpix 0.834 --rise 2.42 --twist 179.45 --z_percentage 0.25
The arguments used in the command above are as follows:
--impose apply real-space helical symmetry
--i input file
--o output file
--cyl_outer_diameter outer diameter of the cylindrical mask
--angpix pixel size in angstroms
--rise helical rise in angstroms
--twist helical twist in degrees
--z_percentage central z-length
F. Model building and validation for alpha-synuclein fibrils
There are many methods for building molecular models. Here, we used PDB 6H6B as a starting point; the model was fit into the EM map using ChimeraX and then one subunit was rebuilt and refined in Coot. The monomer model was subjected to a round of real-space refinement in Phenix. Then, ChimeraX was used to fit additional refined subunits into the map to generate a multimer model. The multimer model was subjected to a final round of real-space refinement in Phenix. We encourage users of this protocol to review tutorials and manuals for ChimeraX, Coot, and Phenix before proceeding with model building [41,42,46,50,51]. During the modeling process, users should use our refined model PDB 9CK3 as a reference. An overview of the entire modeling and validation workflow is provided for reference (Figure 15).
Figure 15. Model building and validation protocol for a-synuclein fibrils. Step-by-step protocol for building, refining, and validating an a-syn fibril molecular model. This protocol uses ChimeraX, Coot, and Phenix in an iterative fashion to improve the molecular model.
1. Download PDB 6H6B by running open 6h6b from the ChimeraX command line [12]. This model covers residues 38–95 of the a-syn protein and contains 10 monomers displaying the amyloid fold.
2. Open the final postprocess.mrc file from step G45 and under Volume viewer setstep size to 1.
3. In ChimeraX, use the Fit tool to place PDB 6H6B into the central region of the postprocess.mrc file. You may need to rotate the model to correctly fit the model into the map.
4. Run the command below from the ChimeraX command line to trim the ends of the map for easier visualization of the central region. Then, continue fitting until the model is well-placed on the map.
view orient; clip front -30 back 30
Note: Clipping can be turned off by running clip offfrom the ChimeraX command line.
5. Remove all but one monomer from the model by running split; delete #1.2–10 from the ChimeraX command line. We will use Coot to build and refine one subunit and add additional subunits later.
6. Save the file as a PDB, ensure that Save relative to model is checked, and select the post-processed map in the drop-down menu.
7. In Coot, go to File > Open Coordinates and select the PDB file saved in step 6. Then, go to File > Open Map and select the post-processed map from section G, step 45.
Note: We encourage new users to review the Coot tutorial to become familiar with the software before proceeding (https://www2.mrc-lmb.cam.ac.uk/personal/pemsley/coot/web/tutorial/tutorial.pdf).
8. On the right-hand side is the modeling toolbar; click Map, click Estimate to set the map weight, and then click Ok.
Note: Click the arrow at the bottom of the modeling toolbar and select Icons and text to add the name of each tool to the modeling toolbar.
9. Go to Refine > All-atom Refine to improve the fit of the map to the model. When the refinement is done, click Accept to save the refined atom positions. The refinement will impose geometry restraints and Ramachandran restraints but will ignore rotamers. We will handle rotamers in Phenix.
Note: Use the mouse scroll wheel to adjust the map contour level as necessary throughout this process.
10. The density generated in this protocol allows for the modeling of additional residues not resolved in PDB 6H6B, so we need to build additional regions of the model. Go to leucine 38, located at the n-terminus; from the modeling toolbar, click Add Terminal Residue and click on leucine 38. This will add an alanine residue to the n-terminus. From the modeling toolbar, click Simple Mutate, and then click on alanine 37. A window will appear listing all the amino acids; click Val (V) to change alanine 37 to a valine. From the modeling toolbar, click Real Space Refine Zone, then click on valine 37 and valine 40. This will refine the region between these two residues and improve the fit of the model to the map. Then, click Accept to save the refined atom positions.
11. Repeat step 10 to add glycine 36 to the n-terminus and lysine 96 and lysine 97 to the c-terminus.
12. Next, build residues 15–22 into a well-resolved island of density located near the n-terminus. Go to the island of density and rotate the density so the fibril core is oriented toward the top of the screen. This will help minimize the number of movements needed to place the strand into place. Go to Calculate > Other Modelling Tools > Place Strand Here, set Estimated number of residues in strand to 8 and click Go. A strand comprised of 8 alanine residues should now appear; use the Real Space Refine Zone tool to improve the fit of the strand into the map.
13. Use the Simple Mutate tool to change the alanine strand to the correct residues (V15, V16, A17, A18, A19, E20, K21, T22). Then, use the Real Space Refine Zone tool to further improve the fit of the strand.
Note: The n-terminus (i.e., island of density) folds back toward the fibril core adjacent to residues 36–44, with residue 15 closest to the fibril core.
14. Click on Display Manager and you will see that there are two molecules: one is the PDB that was imported and the second is the new strand that was created. We need to renumber the residues in the new strand, merge the molecules, and fix the chain ID. Go to Edit > Renumber Residues, under Renumber Residue Range of Molecule select the newly generated strand, under Start Residue select N-terminus, in the Apply Offset box provide an integer value to correct for the difference in residue number for the residue that should be valine 15, then click Apply. For example, if the valine on the n-terminus of the strand is labeled as V40, then the offset should be -25 to set the valine to residue 15. Click on the n-terminus valine of the strand to verify the numbering is correct.
15. To merge the molecules, Go to Edit > Merge Molecules, under Append/Insert Molecule(s) select the strand, and under into Molecule select the original PDB from the drop-down menu. Then, click Merge.
16. Change the chain IDs so both fragments are labeled as chain A. Go to Edit > Change Chain IDs, under Change Chain ID in Molecule select the file that contains both fragments (i.e., the recently merged molecule), under From Chain ID select either chain, under Using Residue Selection select Whole Chain, under To Chain ID set this value toA, then click Apply New Chain ID. Repeat the process if the second fragment is labeled anything other than chain A. The fragments should now be one molecule labeled as chain A with a dotted line showing the missing residues from residues 23 to 35 that are not resolved.
17. Refine the new molecule that spans residues 15–22 and 36–97 (Figure 16). Go to Refine > All-atom Refine and, if the atoms are well-positioned, click Accept. If not, manually adjust misplaced atoms by dragging the atoms into place and then click Accept.
Figure 16. a-synuclein map and model. A cross-section shows the molecular model and the cryo-EM map shows excellent agreement. Side-chain densities are clear and allow for modeling residues V15–T22 and G36–K97. Scale bar, 25 Å.
18. Save the coordinates, go to File > Save Coordinates, under Select Molecule Number to Save select the molecules that were refined in step 17, click Select Filename, and save the file to the desired location.
19. Open Phenix and set up a new project.
Note: We encourage new users to review the Phenix tutorial, specifically the real space refinement tutorial, to become familiar with the software before proceeding (https://phenix-online.org/documentation/reference/real_space_refine.html).
20. Under the cryo-EM section, select the Real-space refinement job. Provide the PDB file from Coot as the model file and the post-processed file as the map file. Set Resolution as determined in the final RELION post-processing job; in this case, the resolution is 2.04 Å. Under the Refinement Settings tab, in addition to the default settings, ensure Use secondary structure restraints andRamachandran restraints is checked, set Nproc to 4, click Rotamers, and under Fit, select outliers and poormap, then click Run. Upon completion, the validation report shows that the model statistics are favorable. The Rotamer outliers (%) will be slightly elevated due to a salt bridge that forms between lysine 80 and glutamic acid 46, causing lysine 80 to be a rotamer outlier that is supported by the data.
21. In ChimeraX, open the refined model and the post-processed map. Open the refined model again; now, two models are available. Select the second model and use the Fit tool to place the second monomer into the opposing protofilament. Repeat the process of opening the refined model and fitting it into a new region of the map. For PDB 9CK3, we built a dodecamer model.
22. Once the desired number of subunits are fitted into the map, run combine from the ChimeraX command line to merge the subunits into one model. The command should provide a unique chain ID to each subunit.
23. Repeat step F6 to save the model relative to the post-processed map.
24. In Phenix, repeat real space refinement as in step 20 with the additional parameter Ncs constraints selected. The final validation report shows excellent model statistics with only lysine 80 as a rotamer outlier, as expected (Table 1). This step can be repeated, if necessary. The model is now ready for structure analysis.
Table 1. Data collection, reconstruction, and model statistics
Protein a-syn (apo) fibrils
PDB ID 9CK3
EMDB ID EMD-45639
Data collection and processing
Voltage (kV) 300
Electron exposure (e-/Å2) 40
Number of frames 40
Nominal defocus range (-mm) 0.5–2.5, 0.25 step
Pixel size (Å) 0.834
Symmetry (beyond helical) C1
Micrographs 5,193
Final particles 129,895
Box size (Å) 300
Box size (pixels) 360
Helical symmetry
rise (Å) 2.42
twist (°) 179.45
Helical Z parameter (%)
masking 25
refinement 25
Map resolution (Å) threshold 0.143
Masked 2.04
Unmasked 2.35
Sharpening B factor (Å2) -43
Structure Refinement
Total atoms 5,748
ligands 0
water 0
RMS, bonds (Å) 0.004
RMS, angles (o) 0.643
Ramachandran favored (%) 98.48
Ramachandran outliers (%) 0.00
Rotamer outliers (%) 2.08
Map-model correlation (masked) 0.90
Model B factors
minimum 5.38
mean 41.28
maximum 88.56
MolProbity score 1.42
Clashscore 3.88
Data analysis
Cryo-EM data processing depends on the sample, data collection instrumentation and parameters used, and computational hardware and software. The workflow presented here should provide users with the necessary details for cryo-EM structure determination of amyloid fibrils. The protocol presented here was used to generate a cryo-EM map and atomic model of in vitro–assembled a-syn fibrils. Atomic models have been deposited in the Protein Data Bank (PDB) under accession 9CK3. Cryo-EM maps, including the final map, half-maps, and mask have been deposited in the Electron Microscopy Data Bank (EMDB) under accession EMD-45639.
Validation of protocol
This protocol or parts of it has been used and validated in the following research articles:
• Dhavale et al. [33]. Structure of alpha-synuclein fibrils derived from human Lewy boy dementia tissue. Nature Communications. https://doi.org/10.1038/s41467-024-46832-5.
• Montemayor et al. [52]. Flagellar Structures from the Bacterium Caulobacter crescentus and Implications for Phage ϕ CbK Predation of Multiflagellin Bacteria. Journal of Bacteriology. https://doi.org/10.1128/jb.00399-20.
• Sanchez et al. [53]. Atomic-level architecture of Caulobacter crescentus flagellar filaments provide evidence for multi-flagellin filament stabilization. bioRxiv. https://doi.org/10.1101/2023.07.10.548443.
• The cryo-EM structure in the manuscript has been validated by our submissions to the PDB (9CK3) and EMDB (EMD-45639). https://doi.org/10.2210/pdb9CK3/pdb.
Limitations
The fibrilization conditions presented here are specific to one form of in vitro–assembled a-syn fibrils. Extensive optimization of protein purification and fibrilization conditions, testing buffer conditions, and incubation parameters may be necessary to generate different in vitro forms. The cryo-EM helical reconstruction methods presented here assume that fibrils are both twisting and of sufficient length to determine the crossover distance for helical twist estimates. There are cases where fibrils may not twist, and thus, this workflow would not be amendable to such samples. Finally, structure determination of patient-derived fibrils is of high interest, but extraction of fibrils from patient tissue is outside of the scope of the work presented here. Though, in theory, the data processing methods presented here should be applicable to these samples.
General notes and troubleshooting
Troubleshooting
The workflow presented here, including sample preparation, NS-TEM, cryo-EM data collection, cryo-EM data processing, and molecular model building, serves as a starting point for individuals new to cryo-EM structural analyses of amyloid proteins. For cryo-EM structure determination, new samples will pose their own unique set of challenges, but by first completing the data processing workflow in section E with the EMPIAR-12229 dataset under accession EMPIAR-12229, new users will be more adept at troubleshooting new issues.
Acknowledgments
This work was supported in part by the University of Wisconsin, Madison, the Department of Biochemistry at the University of Wisconsin, Madison, and public health service grants U24 GM139168 to E.R.W, P41GM136463 to C.M.R, and RF1 NS110436 E.R.W. and C.M.R. from the NIH. J.C.S. was supported in part by the Biotechnology Training Program at the University of Wisconsin, Madison, T32 GM135066, the Steenbock Predoctoral Graduate Fellowship administered by the University of Wisconsin-Madison Department of Biochemistry, and the SciMed Graduate Research Scholars Fellowship with support for this fellowship provided by the Graduate School, part of the Office of Vice Chancellor for Research and Graduate Education at the University of Wisconsin-Madison, with funding from the Wisconsin Alumni Research Foundation and the UW-Madison. C.G.B. was supported by the NIH Ruth L. Kirschstein Fellowship, F32 GM149118, from the NIGMS. We are grateful for the critical feedback, guidance, and support provided by Dr. Bryan Sibert, Dr. Matthew Larson, and Ms. Jennifer Scheuren on cryo-EM data collection, data processing, and use of the cryo-EM HPC cluster. We are grateful to Mr. Owen Warmuth for assistance with the SDS-PAGE gel image. We are grateful for the use of facilities and instrumentation at the Cryo-EM Research Center in the Department of Biochemistry at the University of Wisconsin, Madison. We are grateful for the computational resources supplied through the SBGrid Consortium [34].
Data deposition
The atomic model was deposited in the Protein Data Bank under accession 9CK3. Cryo-EM maps were deposited in the Electron Microscopy Data Bank under accession EMD-45639. The raw micrographs, gain file, and detector MTF file are available on the EMPIAR-12229 database under accession EMPIAR-12229.
Competing interests
The authors declare no competing interests.
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Article Information
Publication history
Received: Aug 18, 2024
Accepted: Nov 28, 2024
Available online: Dec 17, 2024
Published: Feb 5, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Biophysics > Microscopy > Cryogenic microscopy
Biochemistry > Protein > Imaging
Biophysics > Electron cryotomography
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Protocol to Identify Unknown Flanking DNA Using Partially Overlapping Primer-based PCR for Genome Walking
MJ Mengya Jia
DD Dongqin Ding
XL Xiaohua Liu
HL Haixing Li
Published: Vol 15, Iss 3, Feb 5, 2025
DOI: 10.21769/BioProtoc.5172 Views: 64
Reviewed by: Sonali ChaturvediDavid Paul Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in PLOS ONE Mar 2015
Abstract
Genome walking is a popular molecular technique for accessing unknown flanking DNAs, which has been widely used in biology-related fields. Herein, a simple but accurate genome-walking protocol named partially overlapping primer (POP)-based PCR (POP-PCR) is described. This protocol exploits a POP set of three POPs to mediate genome walking. The three POPs have a 10 nt 3' overlap and 15 nt heterologous 5' regions. Therefore, a POP can partially anneal to the previous POP site only at a relatively low temperature (approximately 50 °C). In primary POP-PCR, the low-temperature (25 °C) cycle allows the primary POP to partially anneal to site(s) of an unknown flank and many sites of the genome, synthesizing many single-stranded DNAs. In the subsequent high-temperature (65 °C) cycle, the target single-stranded DNA is converted into double-stranded DNA by the sequence-specific primer, attributed to the presence of this primer complement, while non-target single-stranded DNA cannot become double-stranded because it lacks a binding site for both primers. As a result, only the target DNA is amplified in the remaining 65 °C cycles. In secondary or tertiary POP-PCR, the 50 °C cycle directs the POP to the previous POP site and synthesizes many single-stranded DNAs. However, as in the primary PCR, only the target DNA can be amplified in the subsequent 65 °C cycles. This POP-PCR protocol has many potential applications, such as screening microbes, identifying transgenic sites, or mining new genetic resources.
Key features
• This POP-PCR protocol, built upon the technique developed by Li et al. [1], is universal to genome walking of any species.
• The established protocol relies on the 10 nt 3' overlap among a set of three POPs.
• The first two rounds of POP-PCRs can generally give a positive walking outcome.
Keywords: Genome walking Partially overlapping primer Sequence-specific primer Partial annealing Primary POP-PCR Secondary POP-PCR Tertiary POP-PCR DNA sequencing
Graphical overview
Background
In life science–related areas, genome walking is often employed to obtain unknown genomic DNA flanking known DNA [2–5]. Genome walking has significantly advanced areas such as genetics, biotechnology, and microbiology [6–9]. Many genome-walking methods have been proposed during the past four decades. Although the rationales or experimental steps involved are diverse, these methods can generally be classified into two groups: (i) genome treatment-dependent PCR and (ii) random PCR. The former requires digesting genomic DNA and then ligating the digested DNA with a cassette/adaptor/linker prior to PCR amplification. A random PCR, however, requires just a partial annealing of the walking primer to the unknown flank in the low-temperature (around 30 °C) cycle. A random PCR-based walking protocol has garnered much more attention than other protocols because it omits the time-consuming genome treatment process [10–14].
To date, approximately a dozen techniques of random PCR-based genome walking have been established, such as panhandle (or racket) PCR, thermal asymmetric interlaced PCR, and primer extension refractory PCR [6,12,15–18]. These techniques rely on the random annealing of the arbitrary walking primer to the unknown flanking genomic region, and the subsequent differential amplification between target DNA and non-target DNA. The reason for this differential amplification is that the annealing efficiency of the walking primer is lower than that of the paired sequence-specific primer (SSP). However, these techniques are still comprised of complex PCR steps or unsatisfactory amplification specificity [19–21]. Therefore, researchers have been actively attempting to devise a practical random PCR–based genome-walking technique.
Here, a novel genome-walking protocol of partially overlapping primer-based PCR (POP-PCR) is detailed for efficient genome walking. A POP-PCR set, comprising three rounds of nested amplifications, relies on the 10 nt 3' overlap among the three POPs to overcome non-target amplification. In each round of amplification, the POP has only one chance of partial annealing, as only one 25 °C or 50 °C cycle is performed in this amplification. Thus, only the target product can be amplified, while the non-target product is removed. In addition, unlike other walking methods, POP-PCR uses a unique POP for each round of amplification, which helps reduce non-target background arising from the other POPs. The proposed protocol can be used in many aspects of life science–related areas, like isolating promoters, unveiling new genetic resources, or identifying T-DNA [1,22–24].
Materials and reagents
Biological materials
1. Genomic DNA of rice, given by the Lab of Dr. Xiaojue Peng at Nanchang University
2. Genomic DNA of Pichia pastoris GS115, extracted using Dr. GenTLE (from Yeast) High Recovery kit (Takara, Dalian, China) according to the supplier’s instruction
3. Genomic DNA of Levilactobacillus brevis NCL912 [25–27], extracted using the method described by Li et al. [26]
4. Genomic DNA of human blood, extracted using the method described by Gustafson et al. [28]
Reagents
1. 10× LA PCR buffer (Mg2+ plus) (Takara, catalog number: RR042A)
2. 6× Loading buffer (Takara, catalog number: 9156)
3. LA Taq polymerase (hot-start version) (Takara, catalog number: RR042A)
4. dNTP mixture (Takara, catalog number: RR042A)
5. DL 5,000 DNA marker (Takara, catalog number: 3428Q)
6. 1× TE buffer (Sangon, catalog number: B548106)
7. Agarose (Sangon, catalog number: A620014)
8. 1 M NaOH (Yuanye, catalog number: B28412)
9. 0.5 M EDTA (Solarbio, catalog number: B540625)
10. Green fluorescent nucleic acid dye (10,000×) (Solarbio, catalog number: G8140)
11. Tris (Solarbio, catalog number: T8060)
12. Boric acid (Solarbio, catalog number: B8110)
13. Agarose Gel DNA Purification kit V2.0 (Takara, catalog number: DV805A)
14. Primers (Sangon)
pPOP1: AGTCAGCGTCCAGGTAGTCAGTCTC
sPOP1: TCAGGTCCAAGGTCAAGTCAGTCTC
tPOP1: CTCAGCGTGTTCGTCAGTCAGTCTC
pPOP2: CAGTCAGTCTCAGGTCGTCTCCAGT
sPOP2: AGCAGGTCAGTTACACGTCTCCAGT
tPOP2: TCAGTCAGTCAGTTGCGTCTCCAGT
pPOP3: CGCTTCAGATGGTACAGTGCAGTCA
sPOP3: ACACGATCCCAAGGTAGTGCAGTCA
tPOP3: GTTACTCAGGTCCCAAGTGCAGTCA
pPOP4: GCCTTGAACTGGACCTGATCGACTG
sPOP4: CATGACCGTGCTGAGTGATCGACTG
tPOP4: TGGACTGTGCTACCTTGATCGACTG
gadA-SSP1(5'): CATTTCCATAGGTTGCTCCAAGGTC
gadA-SSP2(5'): ACGTCATCTCAGTTGTTAGCCAACC
gadA-SSP3(5'): AGCCGGTTTGCTTTCAAATGATTCT
gadA-SSP1(3'): TGCGGATACTGATAACAAGACGACA
gadA-SSP2(3'): GGATTGAGAAAGAACGTACGGGTGA
gadA-SSP3(3'): TCCTGCATATCGGTAACGCCCAATC
ALDOA-SSP1(5'): AAATGCTGCAGCCTCCCTCTCACCC
ALDOA-SSP2(5'): AATACCAGAAATGTGCCCTCCCGTG
ALDOA-SSP3(5'): TGAGCTGGCAGGTTGTAGTCTCTGT
ALDOA-SSP1(3'): CCCTCGGACGATTGGACCTAGCTTG
ALDOA-SSP2(3'): GGTCTAACGGTGCCTCTCAGCCTCT
ALDOA-SSP3(3'): TCTGCCCTTCCCCATGGACGTAAGT
malQ-SSP1(5'): CTTCCTGGGTAAGCGTCAGCGTGTG
malQ-SSP2(5'): CAGCTTCGTCGGTAGATTGAACGCT
malQ-SSP3(5'): GGTGGTCAGCAGCCAGCTATATTCG
malQ-SSP1(3'): CGTCATCGCTGTATGGTGATTGGTG
malQ-SSP2(3'): CGGTGTTTACTCCTACAAAGTGCTC
malQ-SSP3(3'): GCTACATTGCCGACAGTAACAGTGC
hyg-SSP1(5'): CGGCAATTTCGATGATGCAGCTTGG
hyg-SSP2(5'): CGGGACTGTCGGGCGTACACAAATC
hyg-SSP3(5'): GACCGATGGCTGTGTAGAAGTACTC
hyg-SSP1(3'): AACTCCCCAATGTCAAGCACTTCCG
hyg-SSP2(3'): GAAACCATCGGCGCAGCTATTTACC
hyg-SSP3(3'): GAAAGCACGAGATTCTTCGCCCTCC
Solutions
1. 2.5× TBE buffer (see Recipes)
2. 0.5× TBE buffer (see Recipes)
3. 100 μM primer (see Recipes)
4. 10 μM primer (see Recipes)
5. 1.5% agarose gel (see Recipes)
Recipes
1. 2.5× TBE buffer
Reagent Final concentration Amount
0.5 M EDTA solution 5 mM 10 mL
Tris 225 mM 27 g
Boric acid 225 mM 13.75 g
ddH2O n/a 950 mL
Total n/a 1,000 mL
Adjust the pH to 8.3 with 1 M NaOH and then replenish the solution to 1,000 mL with ddH2O. This buffer can be stored at room temperature for three months.
2. 0.5× TBE buffer
Reagent Final concentration Amount
2.5× TBE buffer 0.5× 200 mL
ddH2O n/a 800 mL
Total n/a 1,000 mL
This buffer can be stored at room temperature for three months.
3. 100 μM primer
Reagent Final concentration Quantity or Volume
Powdery primer 100 μM n/a
1× TE buffer 1× Volume specified in the sheet of primer synthesis
Total n/a Volume specified in the sheet of primer synthesis
Note: Dilute a portion of the 100 μM primer to prepare 10 μM primer and store the remaining portion at -80 °C.
4. 10 μM primer
Reagent Final concentration Quantity or Volume
100 μM primer 10 μM 1 μL
1× TE buffer 1× 9 μL
Total n/a 10 μL
Note: Prepare extra volume of a 10 μM primer and divide it into multiple 1.5 mL microcentrifuge tubes; then, store the microcentrifuge tubes at -80 °C. Take one tube at a time and store it at -20 °C after use.
5. 1.5% agarose gel
Reagent Final concentration Quantity or Volume
Agarose 1.5% 1.5 g
0.5× TBE buffer 0.5× 100 mL
Green fluorescent nucleic acid dye (10,000×) 1× 10 μL
Total n/a 100 mL
Laboratory supplies
1. 0.2 mL thin-wall PCR tubes (Kirgen, catalog number: KG2311)
2. 10 μL pipette tips (Sangon, catalog number: F600215)
3. 200 μL pipette tips (Sangon, catalog number: F600227)
4. 1,000 μL pipette tips (Sangon, catalog number: F630101)
5. 1.5 mL microcentrifuge tubes (Labselect, catalog number: MCT-001-150)
Equipment
1. PCR apparatus (Applied Biosystems, model: GeneAmp PCR System 2700)
2. Electrophoresis apparatus (Beijing Liuyi, model: DYY-6C)
3. Gel imaging system (Bio-Rad, model: ChemiDoc XRS+)
4. Microcentrifuge (Tiangen, model: TGear)
Software and datasets
1. Oligo 7 software (Molecular Biology Insights, Inc., USA)
2. DNASTAR Lasergene software (DNASTAR, Inc., USA)
Procedure
A. Design of primers
1. Design a POP set of three primers: primary POP (pPOP), secondary POP (sPOP), and tertiary POP (tPOP) (Figure 1).
Figure 1. Interrelationship of the three primers pPOP, sPOP, and tPOP in a POP set. pPOP: Primary partially overlapping primer; sPOP: secondary partially overlapping primer; and tPOP: tertiary partially overlapping primer.
Critical: The sequences of the three POPs are completely arbitrary, comprising a 10 nt of 3' overlap and 15 nt of 5' regions heterologous to each other. Each POP shows a melting temperature of 65–70 °C according to Mazars et al. [29]. A POP itself should avoid forming severe hairpin or dimer structures; meanwhile, it should avoid forming severe dimers with the paired SSP.
Note: Here, the design of the POP1 set (comprising pPOP1, sPOP1, and tPOP1) is provided as an example.
a. Open the Oligo 7 software, click File, then click New Sequence (Figure 2A); type in a 25 nt arbitrary sequence as the initial pPOP1 in the Edit Sequence dialog box (Figure 2B), click Accept/Discard, then click Accept (Figure 2C).
Figure 2. Screenshots showing how to enter an arbitrary DNA sequence in the software. Locations of New Sequence (A), Edit Sequence dialog box (B), and Accept (C) under the File tab.
b. Sequentially click Analyze, Duplex Formation, and Current Oligo (Figure 3A) to check the primer dimer (Figure 3B).
Figure 3. Screenshots showing how to check primer dimer. (A) Locations of Duplex Formation and Current Oligo under the Analyze tab. (B) Predicted primer dimers.
c. Sequentially click Analyze, Hairpin Formation, and Current Oligo (Figure 4A) to check the primer hairpin (Figure 4B).
Figure 4. Screenshots showing how to check primer hairpin. (A) Locations of the Hairpin Formation and Current Oligo under the Analyze tab. (B) Predicted primer hairpins.
Note: Optimize the sequence of this initial pPOP1 as follows if it is unsatisfactory due to forming a severe primer dimer(s) or hairpin(s).
d. Click Edit and Entire Sequence (Figure 5A) to return to the Edit Sequence dialog box (Figure 2B); edit the sequence based on the above analysis results, click Accept/Discard, and click Accept (Figure 2C). Then, minimize this dialog box to show the dialog box shown in Figure 3A.
Figure 5. Screenshots showing how to optimize the sequence of the initial pPOP1. (A) Screenshot showing how to return to the Edit Sequence dialog box. (B) Predicted primer hairpins.
e. Repeat steps A1b and c to evaluate the edited pPOP1, focusing on the formation of primer dimer and hairpin, until a satisfactory pPOP1 (Figure 5B) is obtained.
Note: The pPOP1 shown in Figure 5B is satisfactory, as it only forms an acceptable primer dimer and does not form any hairpin.
f. Fix the 3' part (10 nt) of this satisfactory pPOP1 as the overlap.
g. Attach an arbitrary 15 nt oligo to the 5' end of this overlap, creating an initial sPOP1.
h. Evaluate this sPOP1 using the aforementioned method (namely steps from A1a–e) until a satisfactory sPOP1 is obtained.
i. Design a satisfactory tPOP1 similarly to designing the sPOP1.
Critical: Design more than one POP set, so as to perform parallel POP-PCRs in a genome walking cycle. A severe primer dimer should be avoided between any two POPs in a POP set.
Note: The four POP sets provided by this protocol are universal to genome walking of any species.
2. Select a set of three SSPs (outmost SSP1, middle SSP2, and innermost SSP3) from a known DNA along the direction of 5' to 3'.
Note: Here, the design of gadA-SSP1(5') is provided as an example.
a. Open the Oligo 7 software, click File and then click Open to input the known DNA sequence of gadA gene (Figure 6A).
Figure 6. Screenshots showing how to select a satisfactory gadA-SSP1(5'). (A) Screenshot showing how to input a known DNA sequence. (B) Screenshot showing how to select an oligo with defined length. (C) Predicted primer dimers and hairpin.
b. Click Change and Current Oligo Length to define the length (for example 25 nt) of the oligo to be analyzed; move the mouse to a position of the input sequence (the 25 nt following the mouse is automatically shaded) and then click to select this shaded 25 nt (Figure 6B).
c. Evaluate the selected 25 nt oligo according to the method (Figures 3 and 4) for evaluating the pPOP1.
Note: If the current oligo is unsatisfactory due to forming a severe primer dimer(s) or hairpin(s), re-select and evaluate a new oligo by repeating steps A2a–c until a satisfactory gadA-SSP1(5') (Figure 6C) is obtained.
d. Sequentially design the nested gadA-SSP2(5') and gadA-SSP3(5') like designing the gadA-SSP1(5').
Critical: Each SSP shows a melting temperature of 65–70 °C [29] and should avoid forming severe hairpin or dimer structure.
B. POP-PCR procedure
As shown in Figure 7, a POP-PCR set contains three rounds of nested amplifications. Primary POP-PCR is driven by pPOP and SSP1; secondary POP-PCR is driven by sPOP and SSP2; and tertiary POP-PCR is driven by tPOP and SSP3.
Figure 7. Schematic diagram of POP-PCR. Thin solid line: known sequence; thin dotted line: unknown sequence; arrows: primers; thick lines: primer complements. SSP: sequence-specific primer, pPOP: primary partially overlapping primer, sPOP: secondary partially overlapping primer, and tPOP: tertiary partially overlapping primer.
Note: The primary 25 °C cycle facilitates pPOP partially annealing with a site(s) on the unknown flank and elongating toward the known region to synthesize a target DNA, thus mediating the so-called genome walking. The secondary or tertiary 50 °C cycle facilitates the POP partially annealing with the previous POP site through the 3' overlap. In each PCR, the target single-stranded DNA formed in that 25 °C or 50 °C cycle can be converted into a double strand by the SSP in the following 65 °C cycle and thus can be exponentially amplified by the remaining 65 °C cycles. However, non-target single-stranded DNA formed in that 25 °C or 50 °C cycle cannot be further processed in the following 65 °C cycles, as it lacks a perfect binding site for any primer.
1. Primary POP-PCR
a. Pipette primary POP-PCR components (Table 1) into a 0.2 mL PCR tube.
Table 1. Primary POP-PCR mix
Reagent Final concentration Amount (μL)
Genomic DNA Microbe, 0.2–2 ng/μL; plant or human, 2–20 ng/μL 1
LA Taq polymerase (5 U/μL) 0.05 U/μL 0.5
pPOP (10 μM) 0.2 μM 1
SSP1 (10 μM) 0.2 μM 1
10× LA PCR buffer II (Mg2+ plus) 1× 5
dNTP mixture (2.5 mM each) 0.4 mM each 8
ddH2O n/a 33.5
Total n/a 50
b. Completely mix the components with a pipette.
c. Centrifuge at 1500 g for 10–20 s at 4 °C with a microcentrifuge to gather the mixture.
d. Run the amplification in the PCR apparatus (Table 2).
Table 2. Primary POP-PCR cycling conditions
Step Temp. (°C) Duration Cycle
Initial denaturation 94 1 min 1
Initial denaturation 98 1 min 1
Denaturation 94 30 s 5
Annealing 65 1 min
Extension 72 2 min
Denaturation 94 30 s 1
Annealing 25 1 min
Extension 72 2 min
Denaturation 94 20 s 30
Annealing 65 1 min
Extension 72 2 min
Final extension 72 5 min 1
Hold 4 forever
e. Put the PCR product on ice.
f. Take 1 μL of the product as the template for secondary POP-PCR.
g. Store the remaining product at -20 °C for future assays.
2. Secondary POP-PCR
a. Pipette secondary POP-PCR components (Table 3) into a 0.2 mL PCR tube.
Table 3. Secondary POP-PCR mix
Reagent Final concentration Amount (μL)
Primary PCR product n/a 1
LA Taq polymerase (5 U/μL) 0.05 U/μL 0.5
sPOP (10 μM) 0.2 μM 1
SSP2 (10 μM) 0.2 μM 1
10× LA PCR buffer II (Mg2+ plus) 1× 5
dNTP mixture (2.5 mM each) 0.4 mM each 8
ddH2O n/a 33.5
Total n/a 50
Critical: Dilute primary POP-PCR product 10–1,000 fold if necessary.
b. Completely mix the components with a pipette.
c. Centrifuge at 1500 g for 10–20 s at 4 °C with a microcentrifuge to gather the mixture.
d. Run the amplification in the PCR apparatus (Table 4).
Table 4. Secondary POP-PCR cycling conditions
Step Temp. (°C) Duration Cycle
Denaturation 94 30 s 5
Annealing 65 1 min
Extension 72 2 min
Denaturation 94 30 s 1
Annealing 50 1 min
Extension 72 2 min
Denaturation 94 30 s 30
Annealing 65 1 min
Extension 72 2 min
Final extension 72 5 min 1
Hold 4 forever
e. Put the PCR product on ice.
f. Take 1 μL of the product as the template of tertiary POP-PCR.
g. Store the remaining product at -20 °C for future assays.
3. Tertiary POP-PCR
a. Pipette tertiary POP-PCR components (Table 5) into a 0.2 mL PCR tube.
Table 5. Tertiary POP-PCR mix
Reagent Final concentration Amount (μL)
Secondary PCR product n/a 1
LA Taq polymerase (5 U/μL) 0.05 U/μL 0.5
tPOP (10 μM) 0.2 μM 1
SSP3 (10 μM) 0.2 μM 1
10× LA PCR buffer II (Mg2+ plus) 1× 5
dNTP mixture (2.5 mM each) 0.4 mM each 8
ddH2O n/a 33.5
Total n/a 50
Critical: Dilute secondary POP-PCR product 10–1,000 fold if necessary.
b. Completely mix the components with a pipette.
c. Centrifuge at 1500 g for 10–20 s at 4 °C with a microcentrifuge to gather the mixture.
d. Run the amplification in the PCR apparatus (Table 6).
Table 6. Tertiary fork PCR cycling conditions
Step Temp. (°C) Duration Cycle
Denaturation 94 30 s 5
Annealing 65 1 min
Extension 72 2 min
Denaturation 94 30 s 1
Annealing 50 1 min
Extension 72 2 min
Denaturation 94 30 s 30
Annealing 65 1 min
Extension 72 2 min
Final extension 72 5 min 1
Hold 4 forever
e. Store the PCR product at -20 °C for future assays.
C. Gel electrophoresis
1. Completely mix 5 μL of POP-PCR product and 1 μL of 6× loading buffer.
2. Load the mixture into a 1.5% agarose gel supplemented with 1× green fluorescent nucleic acid dye.
3. Set the electrophoresis apparatus to a voltage of 150 V (the distance between the two electrodes is 30 cm).
4. Check the PCR product using the ChemiDoc XRS+ imaging system after approximately 25 min of electrophoresis (Figure 8).
Figure 8. Genome walking of the gadA locus in Levilactobacillus brevis NCL912 I: walking into the 5' region of gadA, and II: walking into the 3' region of gadA. Each walking comprised four parallel POP-PCR sets: POP1, POP2, POP3, and POP4. The white arrows indicate the target bands. Lanes Ss: secondary PCR products; lanes Ts: tertiary PCR products; lane M1: DL2000 DNA marker; lane M2: λ-Hind III digest DNA marker.
D. Recovery of PCR product
1. Completely mix 40 μL of secondary/tertiary POP-PCR product and 8 μL of 6× loading buffer.
2. Load the mixture into a 1.5% agarose gel supplemented with 1× green fluorescent nucleic acid dye.
3. Set the electrophoresis apparatus to a voltage of 150 V (the distance between the two electrodes is 30 cm).
4. Visualize the PCR product using the ChemiDoc XRS+ imaging system. Subsequently, cut out clear DNA band(s) using a knife.
5. Extract the DNA band(s) from the cut gel using the DiaSpin DNA Gel Extraction kit.
6. Confirm the extracted DNA band(s) with 1.5% agarose gel electrophoresis.
E. DNA sequencing
Sequence the obtained DNA band(s) at Sangon Biotech Co., Ltd (Shanghai, China).
Data analysis
Use the By Clustal W Method function in MegAlign software to analyze the POP-PCR product.
1. Open the MegAlign software, click File and Enter Sequences (Figure 9A) to input a POP-PCR product and the corresponding known DNA segment (Figure 9B).
Figure 9. Screenshots showing how to input DNA sequences. (A) Screenshot of the MegAlign software showing the location of Enter Sequences under the File tab. (B) Input DNA sequences.
2. Click Align and By Clustal W Method (Figure 10A) to output the alignment result (Figure 10B).
Figure 10. Screenshots showing how to align the input sequences. (A) Screenshot of the MegAlign software showing the location of By Clustal W Method under the Align tab. (B) Alignment result.
Note: The walking is considered successful if the SSP3-sided part of the POP-PCR product overlaps the known DNA (an example is shown in Figure 10B).
Validation of protocol
This protocol or parts of it has been used and validated in the following research articles:
• Li et al. [1] Partially Overlapping Primer-Based PCR for Genome Walking. Plos One (Figure 2).
Our POP-PCR has also been validated by several other studies. For instance, Zhang et al. [22] used the POP-PCR to successfully clone the tubulin gene and its promoter region and terminator region in Tribonema minus. Yuan et al. [23] used the POP-PCR to obtain the 3' flanking sequences of the alpha-tubulin gene in Haematococcus pluvialis strains. Wada et al. [24] used the POP-PCR method to identify the insertion site of T-DNA in Arabidopsis T3 6-5. The results indicated that the T-DNA was integrated into Arabidopsis chromosome 3 at position 1334923.
General notes and troubleshooting
General notes
1. The developed POP-PCR comprises three rounds of nested PCR amplifications. However, secondary amplifications can generally release a positive result.
2. Before tertiary POP-PCR, we suggest checking secondary POP-PCR products using agarose gel electrophoresis to know if a clear DNA band(s) appears. If a clear DNA band(s) appears, it is generally not necessary to do tertiary PCR.
3. Like the other PCR-based genome walking schemes, the current POP-PCR scheme also suffers from the issue of multiple DNA bands. However, if multiple DNA bands appear, only the largest band needs to be sequenced.
4. Simultaneously performing parallel POP-PCRs will improve the success and efficiency of genome walking.
5. The current POP-PCR protocol is applicable to genome walking of any species.
Troubleshooting
Problem 1: No clear DNA band(s) is obtained after two or even three rounds of PCRs.
Possible cause: 1) In primary PCR, target amplification is weak; meanwhile, non-target amplification is strong. 2) The annealing temperature (50 °C) of the relaxed cycle in secondary/tertiary PCR is too high.
Solution: 1) Dilute the primary PCR product 10–10,000 times and then use 1 μL of each dilution as the template in the next POP-PCR. Afterward, tertiary PCRs are performed using the secondary PCR products as templates, respectively. 2) Lower the annealing temperature of the relaxed cycle. If still no clear DNA(s) appears in any secondary/tertiary POP-PCR, redesign the SSP set.
Problem 2: A POP-PCR product cannot be directly sequenced.
Possible cause: The non-target background interferes with the sequencing.
Solution: Clone the clear DNA band and then sequence.
Problem 3: Clear DNA band(s) is not a wanted product.
Possible cause: Genomic DNA may be contaminated.
Solution: Re-extract genomic DNA and ensure it is not contaminated; perform PCR amplification in a clean experimental area; ensure that the consumables are sterile.
Acknowledgments
This study was supported by the Major Discipline Academic and Technical Leaders Training Program of Jiangxi Province (grant No. 20225BCJ22023), China. This POP-PCR-based genome walking protocol has been originally described and validated in Plos One [1].
Competing interests
The authors declare no competing interests.
References
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Article Information
Publication history
Received: Sep 13, 2024
Accepted: Dec 2, 2024
Available online: Dec 17, 2024
Published: Feb 5, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Rapid Sampling of Large Quantities of Interstitial Fluid from Human Skin Using Microneedles and a Vacuum-assisted Skin Patch
EW Elizabeth C. Wilkirson
XJ Xue Jiang
PL Peter B. Lillehoj
Published: Vol 15, Iss 3, Feb 5, 2025
DOI: 10.21769/BioProtoc.5173 Views: 62
Reviewed by: Xiyuan LiuPilar Villacampa AlcubierreJessica Lauren Davis
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Original Research Article:
The authors used this protocol in Cell Reports Physical Science Jun 2024
Abstract
Interstitial fluid (ISF) is a promising diagnostic sample due to its extensive biomolecular content while being safer and less invasive to collect than blood. However, existing ISF sampling methods are time-consuming, require specialized equipment, and yield small amounts of fluid (<5 μL). We have recently reported a simple and minimally invasive technique for rapidly sampling larger quantities of dermal ISF using a microneedle (MN) array to generate micropores in the skin from which ISF is extracted using a vacuum-assisted skin patch. Here, we present step-by-step protocols for fabricating the MN array and skin patch, as well as for using them to sample ISF from human skin. Using this technique, an average of 20.8 μL of dermal ISF can be collected within 25 min, which is a ∼6-fold improvement over existing ISF sampling methods. Furthermore, the technique is well-tolerated and does not require the use of expensive or specialized equipment. The ability to collect ample volumes of ISF in a quick and minimally invasive manner will facilitate the analysis of ISF for biomarker discovery and its use for diagnostic testing.
Key features
• Minimally invasive (bloodless and nearly painless) technique for sampling ISF from human skin.
• An average of 20.8 μL of interstitial fluid can be collected within 25 min.
• This technique does not require expensive or specialized equipment or electricity.
• Collected ISF can be analyzed using conventional laboratory-based assays or point-of-care diagnostic tests.
Keywords: Microneedle Interstitial fluid Skin Dermal Biomarker
Graphical overview
Microneedle (MN)-based sampling of dermal interstitial fluid (ISF) using a vacuum-assisted skin patch
Background
The diagnosis, prognosis, and monitoring of many diseases rely on the detection and/or quantification of biomarkers in blood. While blood sampling is a routine medical procedure, it poses risks of infection and can lead to complications, particularly in newborns and individuals with blood clotting disorders [1,2]. In addition, the discomfort associated with blood sampling can deter individuals with blood or needle phobias from getting tested [3,4]. Interstitial fluid (ISF), which is found in the extracellular space in tissues, has garnered much attention as a diagnostic fluid due to its similar biomarker content to blood [5–9]. Prior studies have shown that ISF collected from skin (dermal ISF) is a promising source for biomarkers (e.g., metabolites, proteins, nucleic acids, and exosomes). However, progress in the use of ISF as a diagnostic fluid is hampered by the lack of rapid and minimally invasive methods for collecting ample quantities of fluid [10,11].
Various methods for extracting ISF from skin have been reported, including the creation of suction blisters [7,8,12], microdialysis [13], open-flow microperfusion [14], laser microporation [15], and reverse iontophoresis [16]. While effective, these methods are time-consuming (~1 h), require the use of specialized equipment, and/or involve invasive procedures that can cause skin erythema and dehydration [7]. Microneedle (MN)-based techniques have also been demonstrated for the collection of human ISF, being faster and less invasive [5,12,17,18]. However, the fluid volumes collected from these techniques are too low (~1–5 μL) for biomolecular analysis using conventional diagnostic assays. For example, lateral flow immunochromatographic assays (LFIAs) require at least ~15 μL of sample, western blot requires ~15–60 μL of sample, and enzyme-linked immunosorbent assay (ELISA) requires 50–100 μL of sample.
We have recently reported a simple and minimally invasive method for sampling larger quantities of ISF from human skin [19]. This approach involves the use of a high-density MN array to generate micropores in the skin followed by the attachment of a rigid skin patch and application of mild vacuum pressure using a portable hand pump. The design of the MN array and the parameters associated with the sample collection process (number of MN insertions and the duration of vacuum application) were optimized in our prior study, and this protocol is based on these optimized parameters. The sampling efficiency of this technique was evaluated by collecting dermal ISF from 28 human volunteers, which yielded an average collection volume of 20.8 μL, a ∼6-fold improvement over existing ISF sampling methods. This technique was well-tolerated and reported as being nearly pain-free by all the volunteers. We envision that this protocol will be useful for physicians, scientists, and bioengineers interested in analyzing ISF for biomarkers that can be used for various diagnostic applications, including disease diagnosis and prognosis, as well as for monitoring therapeutic response.
Materials and reagents
Reagents
1. IP-Q photoresin (Nanoscribe, catalog number: 37071000), store in a UV-protected and airtight container in a cool dry environment away from direct sunlight at 4–8 °C; use only under yellow light with λ > 500 nm
2. SU-8 developer (Kayaku Advanced Materials, catalog number: Y020100)
3. Sylgard 184 silicone elastomer kit (Dow, SKU: 4019862)
4. SU-8 2025 photoresist (Kayaku Advanced Materials, catalog number: Y111069-0500L1GL), store in a UV-protected and airtight container in a cool dry environment away from direct sunlight at 4–21 °C; use only under yellow light with λ > 500 nm
5. Dichloro-p-cyclophane (Parylene C dimer) (Specialty Coating Systems, CAS: 28804-46-8)
6. 2-Propanol (IPA) >99.5% ACS (VWR, catalog number: BDH1133-4LG)
7. Deionized water (17.6 MΩ·cm)
8. Liquid nitrogen
9. SU-8 developer (EMD Performance Materials, AZ® Kwik Strip)
Solutions
1. Polydimethylsiloxane (PDMS) (see Recipes)
2. 70% IPA (see Recipes)
Recipes
1. Polydimethylsiloxane (PDMS)
Reagent Final concentration Quantity or Volume
184 silicone elastomer base n/a 40 g
184 silicone elastomer curing agent n/a 4 g
Total n/a 44 g
2. 70% isopropyl alcohol (IPA)
Reagent Final concentration Quantity or Volume
2-Propanol 100% 70 mL
Deionized water n/a 30 mL
Total n/a 100 mL
Laboratory supplies
1. 3 mm thick clear poly(methyl methacrylate) (PMMA) sheet (McMaster Carr, catalog number: 87225K23)
2. 1.5 mm thick clear PMMA sheet (McMaster Carr, catalog number: 8574K51)
3. Polystyrene Petri dish (VWR, catalog number: 89230-472)
4. Plastic mixing cup (Fisher Scientific, catalog number: S04202)
5. Disposable stirring spatula (Millipore Sigma, catalog number: BR759800-500EA)
6. Razor blade (Bates, model: RZ50)
7. Kimwipes (Fisher Scientific, catalog number: 06-666)
8. Microscope glass slide (Fisher Scientific, catalog number: 125444)
9. Micropipette tips [Fisher Scientific, catalog numbers: 02-707-430 (0–200 μL), 02-707-432 (2–20 μL), 02-707-404 (100–1000 μL)]
10. 50 mL conical centrifuge tubes (Fisher Scientific, catalog number: 339653)
11. Double-sided microfluidic tape (3M Company, model: 9972A)
12. Medical-grade pressured-sensitive adhesive tape (Adhesives Research, model: 90106NB)
13. Alcohol prep pad (Fisher Healthcare, catalog number: 22-363-750)
14. Capillary tube, 70 μL (Fisher Scientific, catalog number: 22-260943)
15. Capillary plunger (CoaguSense, catalog number: 03P52-54)
16. Adhesive remover wipes (Smith & Nephew, catalog number: 402300)
17. 0.5 mL protein low-bind microcentrifuge tubes (Eppendorf, catalog number: 4030-8434)
18. 42 mm diameter vacuum cup and hand pump (Hansol Medical Equipment, model: 2018-01-26-0775)
19. Microneedle spring-loaded applicator (Micropoint Technologies)
20. Tweezers (Fisher Scientific, catalog number: 12-000-127)
21. 250 mL glass beaker (Corning Pyrex, catalog number: 1003-250)
22. Nitrile gloves (VWR, catalog number: 76518-336)
Equipment
1. Photonic lithography system (Nanoscribe, model: Photonic Professional GT+)
2. Forced air oven (VWR, catalog number: 89511-410)
3. 6 × 50 mL angle rotor and laboratory centrifuge (VWR, catalog numbers: 76181-202, 76181-190)
4. MiniSpin plus mini centrifuge (Eppendorf, catalog number: 022620207)
5. 50 W, 365 nm UV lamp (SUNUV)
6. Parylene deposition system (Specialty Coating Systems, model: PDS 2010 Labcoater)
7. Digital microscope (Keyence Corporation, model: VHX-7000)
8. CO2 laser cutter (Universal Laser Systems, model: VLS3.75)
9. Dremel MultiPro rotary tool (Dremel, model: 395)
10. Liquid nitrogen dewar (U.S. Solid, model: USS-LNT00001)
11. Nalgene vacuum chamber (Thermo Scientific, catalog number: 5305-0609)
12. Laboport N 816 pump (Fisher Scientific, catalog number: 13-880-31)
13. Portable precision balance (Ohaus, model: SPX2201)
14. Digital timer (Fisher Scientific, catalog number: 06-664-252)
15. Micropipette (Eppendorf, model: 2231300004)
Software and datasets
1. NX Student Edition (Siemens Digital Industries Software, Version 1934, 2020)
2. DeScribe (Nanoscribe, Photonic Professional GT+, 2021)
3. AutoCAD (Autodesk, Student version 2021)
4. Prism version 9.5 (GraphPad Software)
Procedure
A. Fabrication of the MN array master
1. Use NX Student Edition software to create the design for the MN array according to the specifications below.
2. Design a MN with a conical shape having a base diameter of 200 μm and height of 450 μm (Figure S1A).
3. Create a 20 × 20 array of MNs with a needle-to-needle spacing of 400 μm on a 10 × 10 mm square base (thickness = 1 mm) (Figure S1B). See Note 1 for alternative MN design considerations.
4. Save the design as a .stl file.
5. Load the .stl file into Nanoscribe DeScribe software and convert the file to a .gwl file.
6. Load the files into a Photonic Professional GT+ lithography system and print the master using IP-Q resin and a 10× lens, following the manufacturer’s recommended procedure (Figure 1A) (Notes 2–3).
7. After printing, fill one glass beaker with 50 mL of SU-8 developer and another glass beaker with 50 mL of 100% IPA. Use tweezers to gently submerge the master in SU-8 developer for approximately 2 min, then submerge the master in 100% IPA for approximately 2 min.
8. Inspect the master using a microscope to ensure that it was printed correctly and does not have any defects or imperfections (e.g., bubbles, cracks, or broken tips) (Troubleshooting Note 1).
9. Use AutoCAD software to create a 10 × 10 mm square to be used as a substrate for the master.
10. Export the drawing to a laser cutter as a .dwg file.
11. Use the laser cutter to cut the substrate from a 3 mm thick PMMA sheet with double-sided adhesive attached to one side (Note 4).
12. Adhere the substrate to the backside of the master (Figure 1B).
B. Fabrication of the master mold
1. Place the master in the middle of a Petri dish with the MNs facing upward.
2. Combine 40 g of Sylgard 184 elastomer base with 4 g of curing agent in a mixing cup and stir the mixture vigorously for 2 min using a disposable stir rod.
3. Place the PDMS mixture into a vacuum chamber and open the vacuum valve (vacuum gauge level should go past 15 inch/Hg). Degas for 30 min (most of the air bubbles should be removed from the mixture).
4. Pour the degassed PDMS mixture onto the master in the Petri dish. Ensure the master is completely covered (Figure 1C).
5. Place the Petri dish into a vacuum chamber and open the vacuum valve (vacuum gauge level should go past 15 inch/Hg). Degas for 30 min or until all the air bubbles are removed from the mixture.
6. Place the Petri dish in a convection oven preheated at 80 °C for 2 h to cure the PDMS mixture.
7. Remove the Petri dish from the oven and allow the cured PDMS to cool to room temperature.
8. Use a razor blade to carefully cut out the cured PDMS from the Petri dish.
9. Remove the master from the cured PDMS by slowly peeling it out and cut off excess PDMS to create the master mold (Figure 1D).
10. Inspect the mold using a microscope to ensure that it does not contain any defects or imperfections (e.g., bubbles, voids, or uncured PDMS) (Troubleshooting Note 2).
C. Fabrication of the MN array replica
1. Use tweezers to submerge the mold in 70% IPA for 30 min.
2. Remove the mold from the IPA solution and let it air-dry for at least 1 h at room temperature.
3. Place the dried mold on a flat, stable surface and pour SU-8 2025 photoresist into the mold, ensuring the mold cavity is completely filled (Figure 1E).
4. Use tweezers to place the mold into a 50 mL centrifuge tube (Note 5). Ensure the MN tips are oriented toward the bottom of the centrifuge.
5. Centrifuge the tube at 2,500× g for 15 min (Figure 1F).
6. Remove the mold from the tube and place it under a UV lamp for 3 min to cure the SU-8 photoresist (Figure 1G).
7. Remove the cured MN array replica from the mold using gloved hands (Figure 1H).
8. Inspect the replica using a microscope to ensure that it does not contain any defects or imperfections (e.g., missing or cracked tips) (Troubleshooting Note 3).
9. Place the MN array replica into a parylene deposition system. Load the system with 1 g of parylene C dimer and run the coating process following the manufacturer’s recommended procedure (Notes 6–7).
10. Remove the parylene-coated MN array from the deposition system and inspect it using a microscope to ensure that the array is completely coated and the coating is uniform (Figure 1I) (Troubleshooting Note 4). See Note 8 for guidance on batch fabricating MN array replicas.
11. The MN array replica(s) can be stored in a Petri dish (MN tips facing upward) at ambient conditions.
Figure 1. Overview of microneedle (MN) array fabrication procedure. A. The MN array master is fabricated using a Photonic Professional GT lithography system. B. A 3 mm thick PMMA substrate is attached to the backside of the master. C. PDMS is poured over the master and cured in the oven at 80 °C for 2 h. D. The MN master is removed from the PDMS and carefully cut to make the master mold. E. SU-8 photoresist is drop-casted onto the mold. F. Mold is centrifuged at 2,500× g for 15 min. G. SU-8-filled mold is exposed to 365 nm UV light for 3 min to cure the photoresist. H. MN array replica is removed from the mold. I. MN array replica is coated with parylene. This figure is adapted with permission from Jiang et al., 2024.
D. Fabrication of skin patch
1. Use AutoCAD software to create the design for the skin patch according to the specifications below.
2. Design the skin patch as a 42 mm diameter circle with a 2 × 2 array of 11 × 11 mm square cutouts (MN insertion sites) with 2 mm spacing (Figure S2). See Note 1 for alternative skin patch design considerations.
3. Export the drawing to a laser cutter as a .dwg file.
4. Use the laser cutter to cut the patch from a 1.5 mm thick PMMA sheet with double-sided microfluidic tape attached to one side.
5. Use a Dremel rotary tool to smooth the interior edges of the cutouts.
6. Spray the patch with 70% IPA and wipe it clean using a Kimwipe to remove any debris.
7. Use a laser cutter to cut medical-grade adhesive tape with the patch design to generate the skin patch sticker.
E. ISF sample collection from human volunteers
1. Remove the MN array from the Petri dish and rinse it with 70% IPA followed by air drying at room temperature.
2. Wipe the anterior forearm of the volunteer using a fresh alcohol prep pad and allow it to dry (Note 9).
3. Adhere the skin patch sticker to the cleaned forearm.
4. Load the MN array into an MN applicator following the manufacturer’s instructions.
5. Align the MN array over one of the MN insertion sites and place the MN applicator on the sticker (Figure 2A). Press the applicator button to apply the MN array to the skin.
6. Reload the MN applicator with the same MN array following the manufacturer’s instructions. Repeat step E5 two more times for a total of three MN applications per insertion site.
7. Repeat steps E5–6 using the same MN array at the remaining three insertion sites for a total of 12 MN applications (Note 10).
8. Immediately following the completion of all 12 MN applications, start a timer for 3 min (Note 11).
9. While the timer is counting down, remove the backing from the sticker and adhere the skin patch to it, ensuring that the cutouts are aligned and that the patch is firmly attached to the sticker.
10. While the timer is counting down, remove the backing from the skin patch and adhere the vacuum cup to it, ensuring that they are securely attached with no gaps between the cup and patch.
11. After 3 min have passed, attach the hand pump to the vacuum cup and fully pull the handle back once to generate vacuum pressure inside the cup (Figure 2B) (Troubleshooting Note 5).
12. Remove the hand pump from the vacuum cup and maintain vacuum pressure for 20 min (Note 12) (Troubleshooting Note 5).
13. Observe the cutouts of the skin patch for the formation of droplets of ISF on the skin, which should appear ~5–10 min after vacuum application (Figure 2C) (Troubleshooting Notes 6–7).
14. After 20 min have passed, release the vacuum by pulling the pressure release valve on the top of the vacuum cup and detach the vacuum cup from the skin patch (Note 13).
15. Use a capillary tube to collect the extracted ISF from the skin (Figure 2D).
16. Remove the patch from the volunteer’s forearm by slowly peeling one edge off the skin and using an adhesive remover pad to gradually peel away the rest of the patch.
17. Wipe the sampling site using a fresh alcohol prep pad.
Figure 2. Overview of interstitial fluid (ISF) sampling procedure. A. The sticker is adhered to the anterior forearm, followed by microneedle (MN) insertion using the MN applicator. B. The skin patch is attached to the sticker, followed by the attachment of a vacuum cup. Vacuum pressure is generated in the cup using a hand pump. C. The hand pump is removed from the vacuum cup and vacuum pressure is maintained for 20 min. During this time, extracted ISF can be observed on the skin. D. The vacuum cup is removed and the extracted ISF is collected using a capillary tube.
F. Sample processing and storage
1. Transfer the collected ISF sample from the capillary tube into a low-bind microcentrifuge tube using a capillary plunger (Note 14).
2. Incubate the sample for 1 h at room temperature to allow any particulates to sediment.
3. Centrifuge the ISF sample at 6,700× g for 10 min using a microcentrifuge.
4. Use a micropipette to transfer the supernatant to a new low-bind microcentrifuge tube.
5. Snap-freeze the supernatant by immersing the microcentrifuge tube in liquid nitrogen for 5 min.
6. Immediately store the snap-frozen sample at -80 °C until further analysis (Note 15).
Data analysis
Data analysis involved in this protocol includes inspecting the fabricated MN array master and replica using microscopy, examining the collected ISF sample via microscopy, and measuring the volume of the collected ISF sample. Briefly, microscopic inspection of the master, replica, and parylene-coated MN array was performed by imaging at 80× magnification. Microscopic examination of all collected ISF samples was performed by imaging the ISF on a glass microscope slide at 200× magnification. The volume of ISF collected from volunteers was determined by aspirating the ISF sample (after being transferred into a low-bind microcentrifuge tube) using a calibrated 2–20 μL micropipette. The collected volume for each volunteer was determined by averaging two independent sample collections. The mean ISF volume collected by this sampling technique was determined by averaging the volumes of samples collected from 28 volunteers. Means, standard deviations, and statistical analyses were conducted using Prism version 9.5.
Validation of protocol
This protocol has been used and validated in the following research article:
Jiang et al. [19]. Microneedle-based sampling of dermal interstitial fluid using a vacuum-assisted skin patch. Cell Reports Physical Science.
The experimental procedure for fabricating the MN array via centrifugation-assisted replica molding is presented in Video S1, which results in the creation of mechanically robust and sharp MNs (Figure 3A). The strength of the MN array was validated through mechanical testing, as reported by Jiang et al. [19]. Force-displacement curves were generated for MN array replicas with different MN designs and array sizes, including the optimized MN design (base diameter = 200 μm, height = 450 μm) and array size (20 × 20) used in this protocol. This MN array can withstand up to 50 N of compression force, which is 1.5-fold larger than the force required to penetrate human skin, without exhibiting any signs of deformation ([19], Figure S1D). The penetration performance of the MNs was evaluated by inserting the MN array into porcine skin and performing histological analysis on skin sections, which revealed the formation of conical micropores in the epidermis ([19], Figure 2C). Optical micrographs of the MN array after 12 skin insertions revealed that the MNs exhibited no discernible deformation or damage ([19], Figure S3D), validating the ability of the MN array to repeatedly penetrate human skin and generate micropores without fracturing or breaking.
The ISF extraction efficiency was validated by sampling dermal ISF from 28 human volunteers, as reported by Jiang et al., [19]. ISF extracted using this technique was clear to light yellowish in color and generally more viscous than sweat (Figure 3B-D). The average volume of ISF collected from all 28 volunteers was 20.8 ± 19.4 μL (mean ± SD) (Figure 3E). Studies evaluating the safety and tolerability of this technique revealed that it was well tolerated by all volunteers with only minor adverse effects (e.g., skin redness, mild swelling, or slight tenderness localized within the skin patch) that completely resolved within 1 day ([19], Figure S8). Furthermore, volunteers rated the sampling technique as being nearly pain-free ([19], 2024, Figure S7), making it potentially more acceptable to individuals with needle and blood phobias.
Figure 3. Validation of microneedle (MN) array fabrication and interstitial fluid (ISF) sample collection. A. Optical micrograph of the MN array. Scale bar, 1,000 μm. Inset shows a close-up view of a single MN. Scale bar, 100 μm. B–D. Photographs of ISF collected on three different volunteers. Scale bars, 10 mm. E. Mean volumes of dermal ISF collected from the 28 volunteers. Each data point represents the average from two independent sample collections from a single volunteer. The horizontal dotted line represents the average volume of all sample collections, n = 28. Panels B–C are adapted with permission from Jiang et al. [19].
Proteomic analysis of dermal ISF collected from five volunteers using this technique was performed using nanoflow liquid chromatography–tandem mass spectrometry, as reported by Jiang et al. [19]. This analysis resulted in the identification of 2,006 distinct proteins in ISF. Of these proteins identified, 610 are associated with diseases as determined by two online biomarker databases (OncoMX [20] and BIONDA [21]), where 98 are also classified in the NCI Early Detection Research Network biomarker database, and five are approved biomarkers by the US Food and Drug Administration. Dermal ISF samples from COVID-19 vaccinees were analyzed for the presence of SARS-CoV-2 neutralizing antibodies using two commercial SARS-CoV-2 neutralizing antibody tests. SARS-CoV-2 neutralization antibodies were detected in dermal ISF from all vaccinees using both assays ([19], Figure 5C–D). These collective results validate dermal ISF as a source of medically relevant protein biomarkers and demonstrate its utility as a diagnostic fluid.
General notes and troubleshooting
General notes
1. Variations in the design of the MNs and array size can be considered and may prove to be beneficial. For example, a different needle height (e.g., 600 μm, 750 μm) or array size (e.g., 10 × 10) may be beneficial for sampling ISF from other parts of the body with different skin thicknesses and/or topography. Modifications to the design of the skin patch (e.g., smaller cutouts, different cutout layout) may be needed depending on the modified MN array design.
2. Special training and cleanroom environments may be required for the use of the Nanoscribe Photonic Professional GT+ lithography system. Alternative 3D printing platforms with high resolution may be suitable to produce similar results.
3. The standard protocol from the manufacturer’s user guide of this model was used without alterations.
4. Special training may be required for the use of a CO2 laser cutter.
5. A custom centrifuge tube adapter can be utilized to help stabilize and position the MN array mold(s) in the proper orientation during centrifugation. The tube adapter can be customized based on the specific centrifuge tube used and the angle of the tube holder within the centrifuge rotor. A schematic of the 3D-printed tube adapter used in this procedure is shown in Figure S3.
6. Special training and cleanroom environments may be required for the use of the parylene deposition system.
7. The manufacturer’s recommendations were followed for the procedures. The manufacturer-provided ratio is 1 g of parylene C dimer = 1.5 μm of coating.
8. MN arrays can be batch-fabricated by creating multiple molds and performing replica molding processes in parallel. For example, multiple molds can be centrifuged at once, and multiple replicas can be placed in the parylene deposition system for parylene coating.
9. Research activities involving human subjects must be reviewed and approved by an Institutional Review Board.
10. MN arrays should be discarded at the completion of a sample collection procedure and handled as sharps waste.
11. A 3-min wait period between MN insertion and vacuum application is recommended to prevent the collection of blood with ISF. A longer wait period can be used, although it was observed that waiting >5 min resulted in the collection of a negligible amount of fluid.
12. Use caution when removing the hand pump from the vacuum cup to ensure that the pressure inside the vacuum cup is not released by accidentally pulling on the release valve located on top of the cup.
13. The vacuum cup can be reused after sterilization by placing it in boiling water for 30 min.
14. The sample can be examined via microscopy to detect the presence of blood cells, which would indicate blood contamination.
15. In section F (Sample processing and storage), steps 2–6 are optional for applications where the sample is to be analyzed without processing (e.g., rapid diagnostic testing using a lateral flow immunoassay).
Troubleshooting
Problem 1: Defects in the MN array master.
Possible cause: There were issues with resin preparation and/or the 3D printing process.
Solution: Allow the IP-Q resin to come to room temperature before usage. Ensure there are no bubbles when the resin is applied to the lens for 3D printing. Alter the printing specifications according to the manufacturer’s recommendations.
Problem 2: Defects in the master mold.
Possible cause: There are voids and/or bubbles, or PDMS is not fully cured.
Solution: Degas the PDMS for a longer amount of time in the vacuum chamber. Leave the PDMS in the oven for a longer period of time.
Problem 3: Inconsistent formation of MN tips.
Possible cause: SU-8 photoresist spilled out of the mold during centrifugation.
Solution: Place a piece of plastic film over the mold after filling it with SU-8 photoresist. Create a conical centrifuge tube adapter that allows the mold to be oriented at a fixed angle parallel to the bench during centrifugation.
Problem 4: Defects in the parylene coating.
Possible cause: The parylene layer is too thin to evenly coat the MNs.
Solution: Increase the amount of parylene C dimer. Alter the deposition parameters according to the manufacturer’s recommendations.
Problem 5: Vacuum pressure is not generated in the vacuum cup or cannot be maintained for 20 min.
Possible cause: Several factors could affect the vacuum seal, including gaps or air bubbles within the adhesive layer or excessive hair on the skin.
Solution: Remove and replace the sticker and/or the skin patch. Ensure the sticker/skin patch/vacuum cup are securely attached together with no gaps or air bubbles within the adhesive. Adhere the sticker to a location on the anterior forearm with minimal hair.
Problem 6: No ISF is extracted after 10–15 min.
Possible cause: There were issues with the MN application process. Differences in individuals’ skin, such as topology, thickness, or elasticity, can affect the MN penetration depth and micropore size, inhibiting ISF extraction.
Solution: Remove the vacuum cup and repeat the sampling procedure at the same location or at a different location (e.g., the other arm). It is not recommended to repeat the sampling procedure more than twice at the same location during a single sample collection session. Intrasubject and intersubject variability in the amount of ISF collected using this protocol is to be expected.
Problem 7: Blood dots are formed on the skin.
Possible cause: Vacuum is applied too quickly following MN insertion, or the vacuum pressure is too strong.
Solution: Increase the wait time between MN insertion and vacuum application to 4–5 min. Use a weaker vacuum pressure by pulling the pump handle halfway to three-quarters of the way back.
Supplementary information
The following supporting information can be downloaded here.
Figure S1. Designs of the MN and MN array
Figure S2. Design of the skin patch
Figure S3. Design of the centrifuge tube adapter
Video S1. Experimental procedure for fabricating the MN array via centrifugation-assisted replica molding
Acknowledgments
This research was funded by the Wellcome Trust [215826/Z/19/Z]. E.C.W provided material based upon work supported by the National Science Foundation Graduate Research Fellowship [1842494]. This protocol was derived from Jiang et al. [19]. We acknowledge the Shared Equipment Authority of Rice University for the use of the cleanroom facilities to fabricate the MN array master.
Competing interests
X.J. is affiliated with Spear Bio, Inc. E.C.W., X.J., and P.B.L. are co-inventors of a US utility patent application filed by Rice University on the subject of this work.
Ethical considerations
All procedures involving humans were conducted under the guidance and approval from the Rice University Institutional Review Board (IRB-FY2021–147). Criteria for participation was as follows: adults or Rice University students, aged 18 or older, with no blood clotting disorders (including hemophilia or factor II, V, VII, X, or XII deficiencies) or known skin allergies to medical adhesives. Potential participants were provided with informed consent to participate in the study. Participants were explained the entirety of the sample collection process prior to beginning the study. Informed consent of all participating subjects was obtained.
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Article Information
Publication history
Received: Sep 13, 2024
Accepted: Dec 2, 2024
Available online: Dec 13, 2024
Published: Feb 5, 2025
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© 2025 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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Simple and Fail-safe Method to Transform Miniprep Escherichia coli Strain K12 Plasmid DNA Into Viable Agrobacterium tumefaciens EHA105 Cells for Plant Genetic Transformation
BS Beenzu Siamalube
EE Emmanuel Ehinmitan
MN Maina Ngotho
JO Justus Onguso
SR Steven Runo
Published: Vol 15, Iss 1, Jan 5, 2025
DOI: 10.21769/BioProtoc.5174 Views: 370
Reviewed by: Mutinda Sylvia Anonymous reviewer(s)
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Abstract
Agrobacterium-mediated gene transformation method is a vital molecular biology technique employed to develop transgenic plants. Plants are genetically engineered to develop disease-free varieties, knock out unsettling traits for crop improvement, or incorporate an antigenic protein to make the plant a green factory for edible vaccines. The method’s robustness was validated through successful transformations, demonstrating its effectiveness as a standard approach for researchers working in plant biotechnology. It enables the introduction of foreign DNA into plant genomes. Conventionally, plant genetic transformation has relied on time-consuming, costly, and technically demanding procedures, such as electroporation and chimeric viruses or biolistic methods, which usually yield variable transformation efficiencies. This study presents a simple and fail-safe protocol that involves a modified freeze-thaw and heat-shock concoction method. This approach involves a streamlined plasmid miniprep procedure to isolate high-quality plasmid DNA from Escherichia coli K12 strain, followed by a target-specific transfer into A. tumefaciens EHA105 strain. The optimized method minimizes DNA degradation and maximizes uptake by Agrobacterium cells, making it a reproducible and accessible protocol for various genetic engineering applications. The transformation efficiency is consistently high, enhancing plasmid uptake while maintaining cell viability, requiring minimal specialized equipment and reagents. The proposed protocol offers significant advantages, including simplicity, reliability, and cost-effectiveness, positioning it as a valuable alternative to traditional techniques in the field of plant biotechnology.
Key features
• Uses liquid nitrogen as a proxy for freezing.
• Plasmid DNA from competent bacterial cells is extracted using a user-friendly high-copy isolation kit.
• A maximum of five consecutive days is sufficient to complete the procedures.
Keywords: Agrobacterium tumefaciens Plasmid DNA Freeze-thaw method Heat-shock Cell viability Genetic transformation Plant biotechnology Escherichia coli Potato
Graphical overview
Freeze-thaw Agrobacterium transformation method
Background
Agrobacterium tumefaciens is a bacterium found in soil that has a natural ability for transient and stable transfer of foreign DNA into plant cells, including dicotyledonous and monocotyledonous species. This makes it a dominant technique for creating transgenic plants [1]. In the field of plant biotechnology, A. tumefaciens is widely employed to genetically modify plant genomes with desirable traits, such as enhanced nutritional content [2], disease resistance [3], and improved yield, or to vehicle plant-derived edible vaccines [4]. Traditional transformation methodologies using A. tumefaciens include electroporation, direct DNA transfer using chemically competent protocols, and the triparental mating method. Electroporation is a widely used approach, involving the application of an electrical pulse in the bacterial membrane to form pores for penetration of the DNA [5]. Despite being extremely efficient, it needs accurately optimized conditions and specialized equipment, limiting its accessibility for routine use [6]. Chemically competent methods, such as polyethylene glycol (PEG)-mediated transformation, are less sophisticated [7]; however, they present lower transformation efficiency in comparison to electroporation. The triparental mating technique, discovered by Herrera-Estrella et al. in 1983, involves the use of three strains, namely a donor, a helper, and a recipient [8]. It is highly effective yet time-consuming and very labor-intensive.
The freeze-thaw method simplifies the transformation process by eliminating the need for complex electroporation steps or triparental mating setup, making it more adaptable and accessible for various laboratory investigations [9]. The use of miniprep plasmid DNA directly from Escherichia coli reduces turnaround time, allowing for simple and rapid plasmid DNA isolation [10], coupled with uncompromised transformation efficiency [11]. This streamlined approach minimizes procedural errors and contamination chances that may occur when performing transformation experiments with more complex methodologies [12]. Additionally, the protocol is cost-effective, as it does not rely on expensive reagents or equipment; hence, it is easily reproducible and suitable, especially for laboratories with limited resources [13]. Also, it reports a low copy number of integrated transgenes, together with the possibility to transfer larger fragments of DNA. Nevertheless, the protocol is not without its limitations, as the transformation efficiency may be lower than that of optimized electroporation, particularly when dealing with complex gene constructs or larger plasmids [14]. Furthermore, the freeze-thaw method may need additional optimization when applied to different plasmid types or Agrobacterium strains, as variations in DNA uptake capabilities can influence transformation success [15]. Regardless of all these limitations, the protocol remains a practical alternative for routine transformation practices, especially in settings where simplicity and reliability are prioritized over maximal efficiency.
The protocol described in this article has broad utility and is a valuable tool in both basic and applied research across diverse areas of synthetic and microbial biotechnology [16]. It can be employed to introduce plasmids carrying reporter genes, selectable markers, and CRISPR/Cas9 components into A. tumefaciens, facilitating studies on gene expression, gene editing, and functional genomics in plants [17]. Similarly, the method can be extended to develop transgenic plants for biopharmaceutical production, environmental remediation, and biofortification.
Materials and reagents
The equipment, materials, and reagents listed below are appropriate for this protocol. Nonetheless, substitutes sourced elsewhere may be employed if they have shown comparable performance.
Biological materials
1. A. tumefaciens EHA105 strain (strains are preserved at -80 °C in our laboratory)
2. E. coli K12 strain (strains are preserved at -80 °C in our laboratory)
3. Plasmid DNA (stored at -80 °C in our laboratory)
Reagents
1. Lysogeny broth (LB) low salt (Sigma-Aldrich, catalog number: L3397-250G)
2. Agar powder, bacteriological grade (Himedia, catalog number: GRM026-500G)
3. Oligonucleotides (forward: AGGAAACAGCTATGACCATGATTACGAATTC, reverse: ACGTTGTAAAACGACGGCCAGTGCCAAGCTT)
4. OneTaq 2× Master Mix with standard buffer (New England Biolabs, catalog number: M0482)
5. 1 Kb Plus DNA marker (Invitrogen, catalog number: 10787-018)
6. SafeView gel stain (Applied Biological Materials, catalog number: G108)
7. DNA loading dye (Biolabs, catalog number: 10158560)
8. LE agarose (Cleaver Scientific, catalog number: 9012-36-6)
9. TAE buffer (Biological Industries, catalog number: 01-870-1A)
10. Nuclease-free water (BioConcept, catalog number: 3-07F04-H)
11. Rifampicin (GoldBio, catalog number: GB-R-120)
12. Kanamycin sulfate powder (Fisher Scientific, catalog number: BP 906-100)
13. Dimethyl sulfoxide (DMSO) (Sigma-Aldrich, catalog number: D8418)
Solutions
1. LB low-salt liquid medium (see Recipes)
2. LB low-salt agar solid medium (see Recipes)
3. 1% gel (see Recipes)
4. Rifampicin (see Recipes)
5. Kanamycin (see Recipes)
Recipes
1. LB low-salt liquid medium, pH 7.0
Reagent Final concentration Quantity or Volume
Distilled water n/a 100 mL
LB low salt 20 g/L 2 g
2. LB low-salt agar solid medium, pH 7.0
Reagent Final concentration Quantity or Volume
Distilled water n/a 100 mL
LB low salt 20 g/L 2 g
Agar 1.12% (w/v) 1.12 g
3. 1% gel
Reagent Final concentration Quantity or Volume
LE agarose n/a 0.5 g
TAE buffer 1× 50 mL
SafeView gel stain n/a 2 μL
4. Rifampicin
Reagent Final concentration Quantity or Volume
Rifampicin 50 mg/mL 0.5 g
DMSO 99.9% 10 mL
5. Kanamycin
Reagent Final concentration Quantity or Volume
Kanamycin sulfate 50 mg/mL 5 g
Distilled water n/a 100 mL
Laboratory supplies
1. Petri plates (Borosil, catalog number: 3160072)
2. 500 mL culture bottle (Beckman, catalog number: 356011)
3. Powder-free nitrile gloves (Starlab, article number: SG-N-M)
4. Parafilm (Amor, catalog number: PM-996)
5. PCR reaction strips (Simport, catalog number: 330180784)
6. 10 μL pipette tips (AHN Biotechnologie, catalog number: P-214782)
7. 1,000 μL pipette tips (Biologix group, catalog number: MO002462ZA1230-12)
8. P10 micropipette (Eppendorf, catalog number: 4861000-0005)
9. P1000 micropipette (Eppendorf, catalog number: 4861000-0001)
10. 50 mL Falcon tubes (Biologix group, catalog number: 10-9502)
11. 1.5 mL microcentrifuge safe-lock tubes (Eppendorf, catalog number: 022363204)
12. Cell spreaders (ISOLAB, catalog number: CPSA20000002)
13. Isolate II Plasmid DNA kit (Bioline Meridian Biosciences, catalog number: BIO-52056)
14. Laboratory forceps (Thermo Fisher Scientific, catalog number: 112805)
15. Spectrophotometer cuvettes (Thermo Fisher Scientific, catalog number: 14-955-127)
16. Syringe microfilters 0.22 μm (GE Healthcare, catalog number: 289320100)
Equipment
1. Water bath (Cole-Parmer, model: WB-200)
2. Gel doc imaging (UVITEC Cambridge, model: UVIOOC H06)
3. Gel electrophoresis (Cleaver Scientific, model: NANOPAC300P)
4. Clean bench (Haier Biomedical, model: HCB-1600H)
5. Growth chamber (Daihan LabTech, model: LG C-5301)
6. Laboratory freezer (Haier Biomedical, catalog number: 0270501824)
7. Laboratory refrigerator (Haier Biomedical, catalog number: 0270501219A)
8. Thermal cycler (Eppendorf AG, model: 22331 Hamburg)
9. Biological safety cabinet (Haier Biomedical, model: HR1200-IIA2-S)
10. Shaking incubator (Winpact, model: SI-200)
11. Refrigerated microcentrifuge (Eppendorf, catalog number: 5430)
12. 50 mL high-efficient centrifuge (Hermile, catalog number: Z 446 K)
13. BioSpectrometer (Eppendorf, catalog number: 6135K1605057)
14. Laboratory water distiller (Liston, model: A 1210)
15. NanoDrop One (Thermo Fisher Scientific, model: 58595 Intertek)
16. Vertical high-pressure sterilization pot (Labnics, model: NVA-104)
17. Weighing balance (Mettler Toledo, model: PB1302-S/FACT)
18. pH meter (HANNA Instruments, model: H12211)
19. Variable speed vortex mixer (Thermo Fisher Scientific, catalog number: 9501200)
Software and datasets
1. Primer3 (4.1.0, 11/04/2017)
Procedure
A. Plasmid DNA extraction
1. Isolation of plasmid DNA from E. coli K12 strain.
a. Grow E. coli K12 cells containing the desired plasmid.
NOTE: The gene construct in this experiment was cloned into the pCAMBIA1301 plasmid, which contains kanamycin resistance gene for bacterial selection [18] and hygromycin B resistance gene for plant selection [19].
CAUTION: The E. coli cells were grown from colony master plates for 16 h in LB low-salt liquid medium supplemented with 50 mg/L kanamycin at 37 °C and 200 rpm in the shaking incubator (Figure S1).
b. Perform a plasmid miniprep to extract high-copy DNA under the biological safety cabinet (Figure 1).
Figure 1. Isolation of high-copy plasmid DNA from E. coli K12 strain. The ISOLATE II Plasmid Mini kit was used to extract plasmid DNA from the bacterial cells containing the expression cassette of the gene of interest.
B. Agrobacterium setup
1. Preparation of Agrobacterium tumefaciens EHA105 cells.
a. Source for A. tumefaciens cells and test their viability.
b. Make 100 mL of LB low-salt agar solid medium (see Recipes) and sterilize it in a high-pressure sterilizing pot at 121 °C for 20 min.
c. Let it cool to 50 °C, then add 100 μL of rifampicin 50 mg/L and dispense 50 mL in two Petri plates, i.e., 25 mL each (Figure 2b).
NOTE: Agrobacterium tumefaciens strain EHA105 contains rifampicin-resistance gene [20].
d. Add 50 μL of kanamycin 50 mg/L to the remaining 50 mL of the media and dispense it in two separate Petri plates (Figure 2a).
e. Allow the media to jellify and then gently streak the A. tumefaciens colonies using sterile pipette tips.
f. Label and seal the plates with parafilm and incubate for 48 h at 28 °C.
Figure 2. Prepared A. tumefaciens EHA105 cells. (A) No growth of colonies on LB solid media supplemented with rifampicin and kanamycin. The colonies were not expected to grow on this media before transformation, as they are devoid of the plasmid. Hence, the observation of no colony growth confirms the viability of this Agrobacterium strain. (B) Visible growth of incubated A. tumefaciens colony inoculated on LB solid media with rifampicin. Well-grown colonies are observed as a double confirmation that the cells are viable and suitable to proceed with the transformation.
2. Grow A. tumefaciens EHA105 cells to the mid-log phase.
a. Take a fresh, large, and distinct colony from the cells on the plate (Figure 2B), working under a clean bench.
b. Inoculate it in 250 mL of liquid LB media supplemented with only rifampicin in a 500 mL culture bottle.
c. Properly seal the culture bottle and incubate on a shaking incubator set at 300 rpm for 24 h at 28 °C.
d. Check for the optic density (OD600 0.4) using the spectrophotometer.
CAUTION: Blank the spectrophotometer with plain LB liquid medium (without antibiotics) inside a sterile cuvette before loading the sample.
3. Making competent cells.
a. Pellet the cells at 4,500× g for 10 min.
b. Discard the supernatant.
c. Resuspend the pellet in 25 mL of fresh LB liquid media supplemented with rifampicin.
d. Aliquot 250 μL of cells in 1.5 mL Eppendorf tubes.
e. Freeze the tubes in liquid nitrogen for 10 s.
CAUTION: See general note 6. Follow safety guidelines when working with liquid nitrogen.
f. Store the tubes at -80 °C.
C. Bacterial transformation
1. Transformation of E. coli plasmid DNA into A. tumefaciens.
a. Thaw frozen competent cells on dry ice for approximately 3 min.
CAUTION: Do not let the last sliver of ice melt.
b. When the cells are close to complete thawing, place on ice immediately.
c. Split the 250 μL into two tubes, to have 125 μL of cells per tube.
d. Add 5 μL of standard mini-prepared binary plasmid DNA.
CRITICAL: A DNA concentration of 50–100 ng is enough to grow the number of colonies needed.
e. Gently mix by flicking the tube with your finger.
f. For precisely 10 s, freeze the tube in liquid nitrogen with the help of laboratory forceps.
g. Vertically immerse the tubes without the cap, submerging them in the liquid nitrogen.
NOTE: Have the water bath nearby or on the same working bench as the liquid nitrogen container.
2. Use heat-shock to facilitate the uptake of DNA by A. tumefaciens competent cells.
a. Place the tubes in a 37 °C water bath for exactly 5 min.
NOTE: Some tubes may pop out due to the change in temperature; this is expected. Beware of them bursting toward your face.
3. Grow the mixture on a shaker.
a. Lay the tubes horizontally in the shaking incubator.
b. Set the tubes to shake at 300 rpm for 60 min at 28 °C.
NOTE: The cells should have attained OD600 0.4 to ascertain exponential growth.
4. Colony selection and confirmation.
a. Plate 50 μL of the transformed A. tumefaciens cells on a Petri plate containing agar LB media supplemented with rifampicin and kanamycin
b. Grow in a 28 °C incubator for 48 h.
NOTE: Between 20 and 500 colonies are expected (Figure 3).
Figure 3. Colonies of transformed A. tumefaciens. Positive colonies following the successful transformation were grown on LB solid media with rifampicin and kanamycin for 48 h in a 28 °C incubator.
5. Run PCR to confirm successfully transformed colonies.
a. Design primers of pCAMBIA1301 binary vector targeting the gene of interest cloned in the expression cassette using Primer3 software (Table 1).
b. Synthesize the primers and dissolve them to 10μM.
CRITICAL: Make a 10× dilution to get 10μM. For 100 μL of 10μM working solution from the 100μM primer stock, add 10 μL of stock to 90 μL of nuclease-free water.
Table 1. Amplification primer sequences for binary plant vector pCAMBIA1301
pCAMBIA1301
Forward primer AGGAAACAGCTATGACCATGA TTACGAATTC
Reverse primer ACGTTGTAAAACGACGGCCAG TGCCAAGCTT
c. Get 1 μL of a distinct colony as DNA template, add primers (Table 2), and carry out the PCR procedure.
Table 2. PCR reaction master mix with standard buffer
Reagent Amount
OneTaq2× master mix 12.5 μL
DNA template 1 μL
10 μM forward primer 0.5 μL
10 μM reverse primer 0.5 μL
Nuclease-free water 10.5 μL
d. Set the PCR conditions on the thermal cycler as shown in Table 3.
Table 3. Thermocycling conditions for the PCR reaction
Step Temp. (°C) Duration No. of cycles
Initial denaturation 95 30 s 1
Denaturation 95 30 s 35
Annealing 53 1 min
Extension 68 1 min per kb
Final extension 68 5 min 1
Hold 4 ∞ -
e. Prepare 50 mL of 1% LE agarose gel.
f. Heat it in a microwave until it is entirely dissolved (usually takes 1 min).
g. Let it cool down and then add 2 μL of SafeView gel staining solution.
CRITICAL: Be careful not to let the gel solidify before adding SafeView.
h. Mix gently and then pour the mixture into a gel tray with a gel comb to create wells.
i. Load 2 μL of the DNA marker in the first well.
j. Add 1 μL of DNA loading dye to each sample, then load them one after the other in the subsequent wells.
k. Set the gel electrophoresis apparatus to run at 160 V for 15 min.
l. Visualize the gel using the gel doc imaging system to assess band size.
6. Grow positive transformants on selection media.
a. Select one distinct positive colony.
b. Grow it on an LB solid plate containing selection media.
c. Seal the plate with parafilm and incubate at 28 °C for 48 h.
d. The positive colony from the master plate is expected to grow as shown in Figure 4.
NOTE: The bacterial culture is ready for plant genetic transformation.
Figure 4. Positively transformed colony on LB medium supplemented with antibiotics. For plant genetic transformation, use a freshly grown positive colony for better results.
Data analysis
In this protocol, plasmid DNA requires stringent quality control measures and validation assessment. Thus, the first step is undertaken to measure and analyze the concentration and purity of the extracted plasmid DNA from the E. coli cells. This is a vital step in determining the quality of the isolation process. The PCR technique and gel electrophoresis are then performed to verify the size of the gene of interest. In this protocol, we present an analysis of the data collected and provide experimental references that align with Xu et al [21].
Data collected from the transformation experiments typically include the counted number of A. tumefaciens colonies that grew on selective media supplemented with correct antibiotics after the freeze-thaw method. Any notable features such as growth rate and colony morphology were observed and recorded. Only distinct colonies from clear LB solid media plates were included, as an indication of successful eyes-test transformation. Data from plates showing signs of contamination were excluded as well as those that appeared abnormal (possible false positives). Data from samples where the DNA quality was below acceptable standards, i.e., A260/A280 ratio outside a 1.8–2.0 range, were disregarded.
A. Nanodrop analysis of plasmid DNA concentration and purity
Using NanoDrop One, the concentration and purity of the extracted plasmid DNA can be measured to assess the quality of the isolated E. coli K12 cells. The concentration is the amount of plasmid DNA in ng/μL, and a volume between 50 and 100 ng/μL is sufficient. The 260/280 purity ratio indicates protein contamination; it should be between 1.8 and 2.0 for pure DNA. Salt and contaminant presence are depicted by a 260/230 ratio, which should be greater than 2.0 for pure DNA. Results obtained show that the concentration, protein contamination, and salt presence of the double-stranded DNA are 88.8 ng/μL, 1.86, and 2.02, respectively (Figure 5).
Figure 5. E. coli K12 plasmid DNA concentration and purity measured by the Nanodrop spectrophotometer method. Based on the Nanodrop software results and measurement analysis, we can deduce that the concentration of the measured sample is 88.8 ng/μL, while the 260/280 purity ratio is 1.86, and the 260/230 purity ratio is 2.02.
B. PCR verification of the positive transformants
In this experiment, we selected one distinct colony as an important indicator for detecting the presence of the gene of interest. PCR was conducted on the purified sample to amplify the gene sequence; we observed the characteristic band size (Figure 6) using gel electrophoresis, which further showed that the expected size was present in the transformed A. tumefaciens cells.
Figure 6. PCR verification of positive transformants. Lane M: DNA marker. The size of the gene of interest is 1,919 bp, and significant enrichment is visible around 2,000 bp in lanes 1 and 2. This ascertains the presence of the gene of interest in the transformed bacterial cells.
Validation of protocol
Parts of this protocol have been used and validated in the following research article:
• The methods described in this paper are widely accepted and have been used in the fields of plant biotechnology and genetic engineering. The protocol showed positive transgenic efficiency of up to 68.79% in Irish potato tetraploid recipient, Desiree cultivar [21]. Similarly, the procedure recorded a successful transformation of Agrobacterium tumefaciens EHA105 cells with plasmid DNA from E. coli strain K12, resulting in a DNA purity ratio A260/A230 of 2.02 (Figure 5). Furthermore, PCR analysis confirmed the presence of the transgene in the transformed Agrobacterium cells, with a 100% amplification rate (Figure 6).
General notes and troubleshooting
General notes
1. To prevent contamination, ensure that conditions are always sterile throughout the experiment. Autoclave all laboratory supplies, sterilize equipment, and do not use expired reagents. Perform the work within a clean bench or biological safety cabinet.
2. Ensure that the plasmid DNA extracted from E. coli cells is free from contaminants and of high quality. For best results, use freshly isolated plasmid DNA, as degradation can lower transformation efficiency.
3. To attain high competence levels, prepare A. tumefaciens cells appropriately. Confirm that cells are at mid-log phase before making them competent, as older or stationary-phase cells are less likely to take up DNA efficiently.
4. After transformation, use suitable selective media to correctly identify successfully transformed A. tumefaciens colonies. Include the right antibiotics corresponding to the plasmid’s resistance markers.
5. Beware of the plasmid type and size; larger plasmids may need protocol modifications, such as increasing the amount of DNA to be eluted.
6. Handle liquid nitrogen with care as it is extremely cold (-196 °C) and can cause severe frostbite burns. Wear protective gear, use proper storage and handling containers with tight-fitting lids, and secure handles. Use in well-ventilated areas of the laboratory, label containers clearly, and store them upright and away from heat sources. When the experiment is completed, follow safety protocols for the disposal of liquid nitrogen.
Troubleshooting
Problem 1: Low transformation efficiencies.
Possible causes:
- Low-quality or degraded plasmid DNA.
- Improper preparation of competent A. tumefaciens cells (e.g., cells not at the correct growth phase).
- Insufficient DNA concentration or volume during the transformation step.
- Suboptimal heat shock conditions.
Solutions:
- Verify the quality and concentration of plasmid DNA using spectrophotometry.
- Confirm that A. tumefaciens cells are at mid-log phase (OD600 around 0.5–0.8) before making them competent.
- Optimize the amount of plasmid DNA used (generally 50–100 ng is sufficient).
- Adjust transformation conditions (e.g., duration, temperature) to fit the cell type and DNA used.
Problem 2: No growth on selective media.
Possible causes:
- Antibiotic concentration may be too high.
- Incorrect antibiotics or selection markers.
- Inefficient DNA uptake by A. tumefaciens cells.
Solutions:
- Check the antibiotic concentration and confirm it is compatible with the selection of transformed A. tumefaciens.
- Verify that correct antibiotics match the resistance markers present on the plasmid.
- Ensure that competent cells were prepared and stored correctly. Freshly prepared cells generally give better results.
Problem 3: Contamination of cultures.
Possible causes:
- Non-sterile technique during handling.
- Contaminated reagents or equipment.
Solutions:
- Perform all steps under sterile conditions and disinfect tools and surfaces regularly.
- Use fresh, sterile reagents and autoclaved consumables. Periodically check cultures for contaminants.
Problem 4: Inconsistent results.
Possible cause: Variations in cell competence, DNA quality, or environmental parameters.
Solutions:
- Maintain consistent growth conditions for bacterial cultures.
- Standardize procedures for DNA extraction and transformation.
- Repeat experiments to confirm reproducibility and refine any variable steps.
Supplementary information
The following supporting information can be downloaded here:
1. Figure S1. Making competent E. coli cells using CaCl2 and the heat-shock method
Acknowledgments
The authors appreciate the Plant Transformation Laboratory at Kenyatta University for providing space and resources to perform this work. The authors acknowledge the Pan African University Institute for Basic Sciences, Technology and Innovation, sponsored by the African Union Commission.
Competing interests
The authors declare no competing interests.
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Received: Sep 13, 2024
Accepted: Dec 3, 2024
Available online: Dec 18, 2024
Published: Jan 5, 2025
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© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Biological Sciences > Biological techniques > Microbiology techniques
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Quantitative, Dynamic Detection of Neuronal Na+ Transients Using Multi-photon Excitation and Fluorescence Lifetime Imaging (FLIM) in Acute Mouse Brain Slices
SE Sara Eitelmann
KK Karl W. Kafitz
CR Christine R. Rose
JM Jan Meyer
Published: Vol 15, Iss 3, Feb 5, 2025
DOI: 10.21769/BioProtoc.5175 Views: 98
Reviewed by: Alberto RissoneRaniki Kumari Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in The Journal of Neuroscience Jan 2022
Abstract
Fluorescence lifetime imaging microscopy (FLIM) is a highly valuable technique in the fluorescence microscopy toolbox because it is essentially independent of indicator concentrations. Conventional fluorescence microscopy analyzes changes in emission intensity. In contrast, FLIM assesses the fluorescence lifetime, which is defined as the time a fluorophore remains in an excited state before emitting a photon. This principle is advantageous in experiments where fluorophore concentrations are expected to change, e.g., due to changes in cell volume. FLIM, however, requires collecting a substantial number of photons to accurately fit distribution plots, which constrains its ability for dynamic imaging. This limitation has recently been overcome by rapidFLIM, which utilizes ultra-low dead-time photodetectors in conjunction with sophisticated rapid electronics. The resulting reduction in dead-time to the picosecond range greatly enhances the potential for achieving high spatio-temporal resolution. Here, we demonstrate the use of multi-photon-based rapidFLIM with the sodium indicator ION NaTRIUM Green-2 (ING-2) for the quantitative, dynamic determination of Na+ concentrations in neurons in acute rodent brain tissue slices. We describe the loading of the dye into neurons and present a procedure for its calibration in situ. We show that rapidFLIM not only allows the unbiased determination of baseline Na+ concentrations but also allows dynamic imaging of changes in intracellular Na+, e.g., induced by inhibition of cellular ATP production. Overall, rapidFLIM, with its greatly improved signal-to-noise ratio and higher spatio-temporal resolution, will also facilitate dynamic measurements using other FLIM probes, particularly those with a low quantum yield.
Key features
• RapidFLIM of the sodium indicator ING-2 enables the intensity-independent recording of neuronal Na+ transients at unparalleled full frame rates of 0.5–1 Hz.
• RapidFLIM is essentially independent of dye concentrations and therefore not affected by dye bleaching.
• Full in situ calibrations enable the quantification of intracellular Na+ changes at high spatio-temporal resolution.
• RapidFLIM of ING-2 allows unbiased determination of cellular Na+ loading also in conditions of strong cell swelling.
Keywords: ING-2 Sodium Multi-photon imaging Fluorescence lifetime imaging microscopy (FLIM) Hippocampus Neuron Chemical ischemia
Graphical overview
Background
Fluorescence lifetime imaging microscopy (FLIM) has emerged as an important technique for gaining a deeper understanding of ion concentration dynamics inside cells of the central nervous system [1–4]. In contrast to widely used intensity-based imaging, FLIM provides information based on the fluorescence lifetime, which is the duration that a fluorophore remains in an excited state before photon emission [5]. For many chemical indicator dyes, such as the widely used calcium indicator Oregon Green 488 BAPTA-1 [1,6], this excited state lifetime is correlated to ion binding. FLIM therefore enables a direct, quantitative readout of ion concentrations, provided that appropriate calibrations are carried out. Importantly, the fluorescence lifetime is largely independent of the concentration of the indicator dyes. This is particularly advantageous in experimental conditions where fluorophore concentrations change, e.g., due to bleaching or changes in cell volume [7]. However, traditional FLIM is subject to so-called dead-time artifacts, which greatly reduce the number of usable photons [8]. Dead-time artifacts result in the occurrence of seemingly shorter lifetimes when the photon count rates exceed a specific threshold, typically between 1% and 5% of the laser repetition rate. This necessitates the use of long collection times, of up to over one minute, to adequately fit the decay of the photon arrival times [9]. The resulting decrease in temporal resolution restricts the ability of FLIM for fast dynamic imaging.
The development of rapidFLIM, which reduces the dead time from approximately 100 ns to roughly 0.7 ns, has overcome this technical issue, allowing much faster repetition rates [10,11]. This enables the FLIM-based dynamic recording of transient changes in ion concentrations that are characteristic of cells of the nervous system, such as fluctuations in the intracellular Na+ concentration [3]. Na+ is integral to brain function, exerting a profound influence on both neurons and astrocytes through a multitude of cellular mechanisms. Disruptions in Na homeostasis result in altered neuronal excitability and impaired neurotransmission and contribute to the development of neurological disorders [12,13]. FLIM measurements of the intracellular Na+ concentration can be performed using the chemical indicator dye ION NaTRIUM Green-2 (ING-2), which shows an approximately 1–1.5-fold change in fluorescence lifetime upon Na+ binding [3,14,15]. ING-2 offers a relatively broad two-photon cross-section, allowing excitation within the range of 750–1,000 nm, while emission peaks between 500 and 600 nm [14,15]. Recently, we demonstrated for the first time that multi-photon-based, rapidFLIM enables dynamic imaging of Na+ transients in neurons in acute hippocampal tissue slices at so far unprecedented spatio-temporal resolution. Notably, this also allowed unbiased quantitative measurement of changes in neuronal Na+ induced by chemical ischemia, i.e., in conditions of strong cell swelling [3]. The present protocol describes the procedures for rapidFLIM for the quantitative recording of intracellular Na+ with ING-2, including the procedures for in situ calibration.
Materials and reagents
Biological materials
1. Balb/c mice bred by the Animal Care and Use Facility of the Heinrich Heine University Düsseldorf. Alternatively, other suitable rodent models can be used (e.g., Janvier, BALB/cJRj)
Reagents
1. Sodium chloride (NaCl) (Roth, catalog number: 3957)
2. Potassium chloride (KCl) (Roth, catalog number: 6781.1)
3. Calcium chloride dihydrate (CaCl2) (Fluka/Honeywell, catalog number: 31307)
4. Magnesium chloride hexahydrate (MgCl2) (Roth, catalog number: 2189.1)
5. Di-sodium hydrogen phosphate (Na2HPO4) (Roth, catalog number: 4984.1)
6. Sodium hydrogen carbonate (NaHCO3) (Applichem, catalog number: 131965)
7. Glucose (Caelo, catalog number: 2580)
8. 2-[4-(2-Hydroxyethyl)-1-piperazine]ethanesulfonic acid (HEPES) (Fisher, catalog number: BP310-500)
9. Magnesium sulfate (MgSO4) (Sigma-Aldrich, catalog number: 230391)
10. D-Gluconic acid sodium salt (Na-gluconate) (Sigma-Aldrich, catalog number: G9005-500G)
11. Gluconic acid potassium salt (K-gluconate) (Roth, catalog number: 2621)
12. Monensin sodium salt (Alfa Aesar, catalog number: J61669)
13. Potassium iodide (Thermo Fisher, catalog number: A12704.18)
14. Erythrosin B analytical standard, ≥ 98.0% (HPLC) (Erythrosin B) (Sigma-Aldrich, catalog number: 87613)
15. Gramicidin (Sigma-Aldrich, catalog number: G5002)
16. Ouabain octahydrate (Merck, catalog number: 4995)
17. ION NaTRIUM GreenTM-2 acetoxymethyl ester (ING-2 AM) (Mobitec, catalog number: 2011F)
18. PluronicTM F-127 (Pluronic) (Invitrogen, catalog number: P6867)
19. Sulforhodamine 101 (SR101) (Sigma-Aldrich, catalog number: S7635-50MG)
20. Dimethyl sulfoxide (DMSO) (Sigma-Aldrich, catalog number: D8418)
Solutions
1. Artificial cerebrospinal fluid (standard ACSF) (see Recipes)
2. Modified ACSF for preparation of brain slices (preparation ACSF) (see Recipes)
3. HEPES-based ACSF (see Recipes)
4. ACSF for calibration (see Recipes)
5. SR101 stock solution (see Recipes)
6. ING-2 AM stock solution (see Recipes)
7. Bolus-loading solution (see Recipes)
8. Instrument response function solution (IRF solution) (see Recipes)
Recipes
1. Standard ACSF
Reagent Final concentration in double-distilled water
NaCl 130 mM
KCl 2.5 mM
CaCl2 2 mM
MgCl2 1 mM
NaH2PO4 1.25 mM
NaHCO3 26 mM
Glucose 10 mM
Adjust pH to 7.4 by bubbling with carbogen (95% O2 and 5% CO2).
The recommended overall volume is 1,000 mL.
2. Preparation ACSF
Reagent Final concentration in double-distilled water
NaCl 130 mM
KCl 2.5 mM
CaCl2 0.5 mM
MgCl2 6 mM
NaH2PO4 1.25 mM
NaHCO3 26 mM
Glucose 10 mM
Adjust pH to 7.4 by bubbling with carbogen (95% O2 and 5% CO2).
The recommended overall volume is 1,000 mL.
3. HEPES-based ACSF
Reagent Final concentration in double-distilled water
NaCl 125 mM
KCl 3 mM
CaCl2 2 mM
MgSO4 2 mM
NaH2PO4 1.25 mM
HEPES 25 mM
Glucose 10 mM
Adjust pH to 7.4 by titration with NaOH.
The recommended overall volume is 100 mL.
4. ACSF for calibration
Note: Prepare calibration solutions with different sodium concentrations (e.g., 0, 10, 50, and 100 mM) by adjusting the concentration of K+, maintaining osmolarity. Solutions need to be prepared freshly every day.
Reagent Final concentration in double-distilled water
Na+ + K+ 150 mM
Cl- 30 mM
Gluconic acid 120 mM
HEPES 10 mM
MgSO4 2 mM
CaCl2 2 mM
Gramicidin 3 μM
Monensin 10 μM
Ouabain 1 mM
Adjust pH to 7.4 by titration with KOH.
The recommended overall volume is 500 mL for each.
5. SR101 stock solution
Reagent Final concentration Quantity or Volume
SR101 1.5 mM -
Double-distilled H2O - 1 mL
6. ING-2 AM stock solution
Reagent Final concentration Quantity or Volume
ING-2 AM 1 mM -
Pluronic 20 mg/mL -
DMSO - 46.12 μL
7. Bolus-loading solution
Reagent Final concentration Quantity or Volume
ING-2 AM stock solution 125 μM 2 μL
HEPES-based ACSF - 14 μL
8. IRF Solution
Note: Dissolve potassium iodide in H2O to saturation. Then, add 0.5 mg of Erythrosin B per 5 mL of H2O.
Reagent Final concentration Quantity or Volume
Potassium iodide Until saturation -
Double-distilled H2O - 5 mL
Erythrosin B - 0.5 mg
Laboratory supplies
1. Cyanoacrylate glue (e.g., Superflex gel, UHU)
2. Razor blades (e.g., razor blades classic, Wilkinson sword)
3. Scalpel blades (e.g., Heinz Herenz)
4. Filter paper (e.g., plain disc filter paper, Lab Logistics Group GmbH)
Equipment
1. Microtome (e.g., Campden Instruments, model: Campden 7000smz-2)
2. Micropipette puller (e.g., Narishige, model: PC-100)
3. Micromanipulator (e.g., Luigs & Neumann, model: LN Junior)
4. Pressure application device for dye injection (e.g., NPI Electronic GmbH, model: PDES)
5. Standard harp slice grids (e.g., ALA Scientific, model: HSG-5A or Warner SHD 41/10)
6. Photon count rate-optimized and custom-modified multiphoton laser imaging system (Figure 1) (a) with a high-intensity pulsed laser source (b) and time-correlated single photon counting (TCSPC) unit for FLIM data acquisition (c), e.g.,
a. Nikon A1MP (Nikon Europe) (Figure 1, steps 1–3)
b. MaiTai DeepSee (Newport Spectra-Physics GmbH, MKS) (Figure 1, step 4)
c. Multiharp 160, PicoQuant GmbH (Figure 1, step 3–4)
Custom modifications mainly include hardware and optical adaptations in the microscope, the NDD stage, and the connection between the microscope and the FLIM detector unit to prevent the reduction of the emission signal intensity going to the FLIM detection unit. These kinds of modifications require the help of both mechanical and electronic workshops and the support of the microscope manufacturer.
Figure 1. Multiphoton microscope setup. 1. Front view of the Nikon A1R MP with system control table. All software necessary for acquisition is started on the adjacent workstations and visible on the respective monitors. 2. Close-up of the microscope stage, with the slice chamber and micromanipulators present in the middle. 3. Backside of the Nikon A1R MP with customized emission port. FLIM unit and incident optical unit are visible in the middle. 4. Simplified scheme of the beam path. In the case of the A1R MP, the excitation beam is directed from the laser to the incident box and optical unit, where it is guided to the scan head of the microscope. After exciting the probe, the emission beam is directed to the FLIM detectors with as few protrusions (e.g., mirrors and filters) as possible, for maximal photon collection efficacy.
Software and datasets
1. NIS Elements Advanced Research (5.21, 08/28/2024, Nikon)
2. SymPhoTime 64 (2.8, 08/28/2024, PicoQuant)
3. OriginPro® (2021, 08/28/2024, OriginLab)
4. Microsoft Excel (Office 2021, 08/28/2024, Microsoft Corporation)
5. Affinity Designer [1.10, 08/28/2024, Serif (Europe) Ltd.]
Procedure
A. Preparation of acute hippocampal brain slices
1. To obtain acute brain slices, mice older than postnatal day 14 are anesthetized with CO2 and subsequently rapidly euthanized by decapitation (for a detailed explanation of animal policies, see also the section Ethical considerations) (Figure 2, step 1). The procedure can also be used for adult animals or animals younger than P14. Note, however, that different protocols for anesthesia may apply.
2. Following decapitation, remove the brain rapidly and place it in a Petri dish containing ice-cold preparation ACSF. To maintain a pH of 7.4, ACSF must be continuously bubbled with carbogen (95% O2 and 5% CO2) (Figure 2, step 2).
3. After the separation of the hemispheres, make an additional incision in a parasagittal orientation (approximately 1.5 mm thick). Affix the tissue block containing the hippocampus with glue to the cutting stage of the microtome at the parasagittal incision. Place the cutting stage into the microtome and cool (ideally with a non-dissolving cooling element) (Figure 2, step 3).
4. Prepare hippocampal slices with a thickness of 250 μm, keeping the tissue constantly submerged in ice-cold preparation ACSF (Figure 2, step 4).
5. Transfer the slices very gently to ACSF containing 1 μM SR101 at 34 °C for 20 min, during which they will take up the astrocyte marker. Following this, incubate the slices at 34 °C for a further 10 min in standard ACSF. Slices can be used for experiments for approximately 5–7 h if kept at room temperature (Figure 2, step 5).
Figure 2. Protocol for acute brain slice preparation. 1. Brain isolation starts by cutting along the illustrated dashed lines. Note: It is necessary to work fast to minimize the time period in which the brain is not submerged in ice-cold ACSF (see below). Always use maximum care to prevent damage to the tissue probes. 2. Excess tissue is removed with additional trimming cuts as depicted by dashed lines. 3. Parasagittal trimming is achieved by cutting a single hemisphere at a 45° angle. 4. The hemisphere is glued to the vibratome cutting stage and placed in the buffer tray filled with ice-cold, carbonated modified ACSF. 5. Freshly cut slices are placed onto a mesh positioned in a beaker filled with modified, ice-cold ACSF. 6. Just before starting experiments, slices are positioned in the experimental chamber and secured with a fine grid. 7. Bolus-loading of hippocampal CA1 region.
B. Bolus-loading of ING-2 into hippocampal brain slices
Note: This technique was initially developed for in vivo loading of dyes into brain cells [16]. It has since been adapted for use with acute brain slices and is employed by numerous research laboratories, including our own.
1. Prepare pipettes for dye injection (tip diameter of approximately 1 μm and a resistance of 1–3 MΩ) from fire-polished borosilicate glass capillaries using a standard micropipette puller.
2. Place a hippocampal brain slice into a microscope chamber and fix it with a “harp” (e.g., ALA Scientific HSG-5A) (Figure 2, step 6). After placing the chamber in your imaging system, constantly perfuse the brain slice with standard ACSF (typical perfusion rate is 2 mL/min).
3. Load an injection pipette with bolus-loading solution (see Recipes) and attach the pipette to the pressure application device.
4. Position the pipette with care and the help of a micromanipulator into your region of choice in the tissue (Figure 2, step 7). Dye injection should be close to the cellular region you are interested in, usually in the same field of view. For loading of somata of neurons in the CA1 pyramidal area, the dye injection pipette should be positioned, e.g., in the stratum radiatum, just below the CA1 pyramidal layer (Figure 3, step 1). Apply ~1.5 psi for ~5 s to inject the dye into the tissue. This process should be repeated until the entire field of view is covered (Figure 3, step 1).
5. Wait for 30–45 min to engage intracellular de-esterification and wash off excess dye from the extracellular space, constantly perfusing the brain slice with standard ACSF.
Figure 3. Calibration process. 1. Inject ING-2 at the region of interest. Here, a fluorescence lifetime image of the CA1 region of a hippocampal tissue slice is shown. Pipettes and approximate areas, dye-loaded by one injection, are indicated in white. 2. Perfuse the slice with Na+-free saline to wash out extracellular Na+. 3. Perfuse with 0 Na+ calibration solution containing ionophores to equilibrate intra- and extracellular Na+. 4. Continue perfusion with calibration solutions at different Na+ concentrations. 5. Plot the resulting lifetimes against Na+ using appropriate software (e.g., Origin Pro). 6. Fit the resulting data to determine the apparent KD(app) and dynamic range of the dye. Modified and taken from [3].
C. Fluorescence lifetime imaging microscopy of ING-2 and multi-photon excitation
1. Choose suitable parameters for imaging for your given imaging system and the specific dye used. With multiphoton excitation, ING-2 can be excited over a broad wavelength spectrum from 740 to 1,100 nm. Typical parameters for ING-2 used in our experiments are an excitation wavelength of 840 nm, an image size of 512 × 512 pixels, and an imaging frequency of 1 Hz.
Note 1: The whole setup should be tuned for maximum photon efficacy. The more photons that can be recorded, the better the spatio-temporal resolution will be. Temporal resolution is always relative, as the recorded frames can be binned afterward. However, a minimum number of photons (for Na+-sensitive dyes 15–20 photons per pixel and frame) need to be recorded for proper lifetime calculation.
Note 2: The setup process before every experiment is crucial to obtain high-quality results. Make sure that the laser power under the objective does not exceed 5 mW. Use appropriate emission filters and dichroic mirrors to ensure optimal detection conditions. For ING-2, emission can be collected above 560 nm wavelength.
2. Measure the instrument response function (IRF) of your microscope with the IRF solution (see Recipes) using comparable photon counts as during experimental conditions (e.g., same laser power).
3. Select your region of interest in your tissue preparation.
Critical: Make sure that your tissue sample is healthy and that many viable neurons are present. Injured neurons are usually characterized by their swollen cell bodies. Moreover, dendrites may be distorted and not be clearly visible.
4. Record an image (e.g., an average of 30 frames) in standard ACSF to check initial conditions.
5. Start a time series in standard ACSF and record fluorescence lifetime in baseline conditions (e.g., for 30 s).
6. Next, record a time series performing your manipulation of choice to test if this results in a change in fluorescence lifetimes (and intracellular Na+). As a positive control, bath application of glutamate (e.g., 1 mM glutamate for 10 s) causes reliable and prominent intracellular Na+ signals in rodent hippocampal slices [17]. Moreover, bath application of inhibitors of cellular ATP production induces long-lasting and large increases in intracellular Na+ in the same preparation [2,3,18]. Alternatively, substances can be focally applied to a soma or a dendrite. This is usually done with a fine glass pipette (comparable to the ones used in section B for bolus application but with a smaller tip diameter) that is filled with the substance of choice (e.g., 1 mM glutamate). The application duration can be a lot shorter in comparison to bath applications and is usually below 1 s [3,19,20].
7. Check if the changes induced by the manipulation are reversible, i.e., if fluorescence lifetimes recover to the initial baseline levels. After the lifetimes have returned to initial values for at least 30 frames, stop the time series.
8. Take another image (e.g., again an average of 30 frames) to compare starting and finishing conditions in standard ACSF.
D. Calibrating changes in lifetime to changes in Na+
Note: The general calibration routine for Na+-sensitive indicator dyes was described in detail by Rose and Ransom [21] for cultured astrocytes. It was later adapted for acute hippocampal brain slices and widefield and confocal as well as multi-photon microscopy [3,22,23]. This protocol was first used for FLIM by Meyer et al. [2].
1. Prepare calibration solutions with different Na+ concentrations, ranging from unsaturated (0 mM Na+) to saturated (>100 mM Na+) concentrations.
Caution: Select a frame rate and laser power, which enables the collection of sufficient photons under all conditions.
2. Start perfusing a dye-loaded tissue slice (Figure 3, step 1) with standard ACSF and then switch to 0 mM Na+ solution without the ionophores and ouabain to wash out Na+ from the tissue (Figure 3, step 2).
3. Record the resulting changes in fluorescence lifetime.
4. Then, perfuse the slice with 0 mM Na+ solution containing the ionophores and ouabain to promote washout of Na+ from the cells and to enable equilibration of Na+ between intra- and extracellular space (Figure 3, step 3).
5. Switch to calibration salines containing different Na+ concentrations (e.g., 10, 20, 30, 50, and 100 mM), again recording changes in fluorescence lifetime resulting from the changes in Na+ (Figure 3, step 4).
Note: Always wait before switching to the next concentration until a stable plateau has been reached. You might need to adjust the focus of your microscope during the calibration process.
6. Plot the changes in lifetime against Na+ and use a Michaelis–Menten equation to fit the data points and to determine the apparent Kd and the dynamic range/saturation of the dye used (Figure 3, step 5).
7. The resulting fit will also allow you to directly convert fluorescence lifetime values obtained during different experiments into Na+ concentrations (Figure 3, step 6).
Data analysis
1. Start by loading an image into your analysis software.
2. Calculate the IRF using the measurement of the IRF solution (see step C2).
3. Select the region of interest you would like to analyze (e.g., a single, neuronal cell body).
4. Determine the lifetime distribution from the chosen region of interest.
5. Fit a mono- or multi-exponential decay to your measured fluorescence lifetime distribution.
Note: Finding optimal fitting parameters can be a tedious task. Decays can be fitted, mono-, bi-, or multi-exponential depending on the used indicators. When fitting with more than one exponent, using fixed exponents can increase the signal-to-noise ratio.
6. Export the fitted results for data visualization and basic analysis in a suitable program (e.g., Excel, Origin Pro).
7. Use an appropriate program for the visualization of the data in the form of figures (e.g., Affinity Designer).
Validation of protocol
This protocol or parts of it has been used and validated in the following research articles:
• Meyer et al. [2]. Quantitative Determination of Cellular [Na+] By Fluorescence Lifetime Imaging With CoroNaGreen. Journal of General Physiology.
• Meyer et al. [3]. Rapid Fluorescence Lifetime Imaging Reveals That TRPV4 Channels Promote Dysregulation of Neuronal Na+ in Ischemia. Journal of Neuroscience.
These two publications illustrate and describe the above-mentioned in situ calibrations in detail, also providing approaches for dye calibration in vitro. Meyer et al. [3] described rapidFLIM in rodent brain tissue slices, also demonstrating that this technique enables reliable, unbiased determination of baseline Na+ concentrations in CA1 pyramidal neurons. Moreover, Meyer et al. [3] presented different manipulations to induce reversible changes in neuronal Na+ and their recording by rapidFLIM. Figure 4 shows that inhibition of cellular metabolism by perfusing tissue slices with glucose-free saline containing inhibitors of glycolysis and oxidative phosphorylation results in large increases in Na+ in CA1 pyramidal neurons.
Figure 4. Quantitative, dynamic measurement of Na+ loading in hippocampal neurons using multi-photon excitation and rapid FLIM with ING-2. A. Color-coded fluorescence lifetime images (30 s temporal binning) of the intracellular Na+ depicting the CA1 pyramidal cell layer of an ING-2-loaded acute hippocampal tissue slice. Right: color code. Top left: image showing intracellular Na+ at baseline conditions; middle: change in Na+ upon mild chemical ischemia (2 min of perfusion with 2 mM NaN3 and 5 mM 2-Desoxyglucose); right: recovery taken ~20 min after starting perfusion with inhibitors. Bottom: similar illustration, demonstrating the neuronal Na+ loading upon moderate chemical ischemia (same solution as before but with 5 min of application). B. Changes in intracellular Na+ induced by mild (2 min; left) and moderate (5 min; right) chemical ischemia (indicated by the gray boxes) reported by rapidFLIM in two different experiments. Gray traces: individual cells; black traces: averages of all neurons analyzed in one particular experiment (n = 35 and 40, respectively). Note that data points were constrained to the minimum/maximum value of the calibration (0 and 150 mM Na+). Images modified and taken from [3].
General notes and troubleshooting
General notes
1. All solutions for experiments and preparation should be prepared freshly on the day of use. Standard and preparation ACSF need to be bubbled with carbogen for at least 30 min before use to ensure a correct pH of 7.4.
2. Acute brain slices need to be treated with utmost care. Bending and other mechanical stress result in damaged cells.
3. As optimizing the experimental setup is critical to obtain optimal photon counts, it is recommended to test imaging parameters in an in vitro environment first before using acute brain slices.
4. Most ion-sensitive dyes are diluted in 20% Pluronic/DMSO. Pluronic is sensitive to freeze and unfreeze cycles, which therefore should be avoided.
5. The total number of collected photons directly correlates to the quality of your measurements. Temporal resolution does directly increase with an increased number of collected photons. The measurement setup should therefore be tweaked for maximum photon collection. This can be achieved by using as few mirrors and filters as possible and using the highest possible grade of optical components.
6. Many manufacturers offer dye spectra and calibration curves obtained in vitro (in the cuvette). Note that most chemical ion indicators change their properties when loaded inside cells. Therefore, in situ calibration is highly recommended.
Troubleshooting
Problem 1: Photon counts are generally too low.
Possible cause: Something might be blocking the light path.
Solution: Check the microscope for any possible obstructions or optical deficits (c.f. problem 4).
Problem 2: Lifetimes are generally too short.
Possible cause: You might be running into photon pileup.
Solution: Modern detectors with lower dead times show nearly no photon pileup effects. Either change detectors or reduce laser power to more adequate count rates.
Problem 3: Staining is too weak.
Possible cause 1: Your staining solution might be old, or ACSF may have been sucked up into the tip of the pipette.
Solution: Prepare a fresh staining solution and control the pre-pressure of your application device.
Possible cause 2: Your preparation is not viable; slices may contain many dead cells.
Solutions: Try to minimize cell damage by treating the tissue with utmost care. Re-adjust your vibratome to minimize vertical blade movements. Pay attention to reducing preparation durations as much as possible.
Problem 4: The photon counts are insufficient for calculating a lifetime, and the statistical fit is inadequate.
Possible cause: You might not have enough photons per frame.
Solutions: Either increase laser power or increase the number of binned frames. Try to optimize the optical configuration of your microscope system by decreasing the number of optical components and increasing the optical quality of the components.
Problem 5: Lifetimes are different than expected and/or not changing upon stimulation.
Possible cause: Crosstalk of autofluorescence or a second fluorescent dye.
Solution: If you are using a second dye for cell identification, check for possible crosstalk and bleed-through. Also, check the autofluorescence of your specimen for possible lifetime artifacts. Crosstalk needs to be eliminated completely to be able to get a proper and stable lifetime readout.
Acknowledgments
The work at the Heinrich Heine University was supported by the German Research Foundation, Deutsche Forschungsgemeinschaft (FOR2795, Synapses under Stress, Ro2327/13-2 to C.R.R.) and by the Federal Ministry of Education and Research (BMBF), Germany (Project SynGluCross to C.R.R.). We thank Simone Durry and Claudia Roderigo, Institute of Neurobiology for expert technical assistance. This protocol was adapted and modified from Meyer et al. [3].
Competing interests
The authors declare no competing interests.
Ethical considerations
Prior to initiating experiments, it is imperative to confirm that all necessary permissions and approvals have been obtained and that experiments are conducted in compliance with all pertinent institutional and national guidelines and regulations. The procedures described here are in strict accordance with the institutional guidelines of the Heinrich Heine University Düsseldorf, Germany, as well as the European Community Council Directive (86/609/EEC). All experiments described here were communicated to and approved by the Animal Welfare Office at the Animal Care and Use Facility of the Heinrich Heine University Düsseldorf, Germany. The relevant institutional act number is O52/05. In accordance with the German Animal Welfare Act (Tierschutzgesetz, Articles 4 and 7), no additional approval was necessary for the post-mortem removal of brain tissue.
References
Zheng, K., Bard, L., Reynolds, J. P., King, C., Jensen, T. P., Gourine, A. V. and Rusakov, D. A. (2015). Time-Resolved Imaging Reveals Heterogeneous Landscapes of Nanomolar Ca2+ in Neurons and Astroglia. Neuron. 88(2): 277–288.
Meyer, J., Untiet, V., Fahlke, C., Gensch, T. and Rose, C. R. (2019). Quantitative determination of cellular [Na+] by fluorescence lifetime imaging with CoroNaGreen. J Gen Physiol. 151(11): 1319–1331.
Meyer, J., Gerkau, N. J., Kafitz, K. W., Patting, M., Jolmes, F., Henneberger, C. and Rose, C. R. (2022). Rapid Fluorescence Lifetime Imaging Reveals That TRPV4 Channels Promote Dysregulation of Neuronal Na+ in Ischemia. J Neurosci. 42(4): 552–566.
Untiet, V., Kovermann, P., Gerkau, N. J., Gensch, T., Rose, C. R. and Fahlke, C. (2017). Glutamate transporter-associated anion channels adjust intracellular chloride concentrations during glial maturation. Glia. 65(2): 388–400.
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Rungta, R. L., Choi, H. B., Tyson, J. R., Malik, A., Dissing-Olesen, L., Lin, P. J., Cain, S. M., Cullis, P. R., Snutch, T. P. and MacVicar, B. A. (2015). The cellular mechanisms of neuronal swelling underlying cytotoxic edema. Cell. 161(3): 610–621.
Becker, W. (2015). Advanced Time-Correlated Single Photon Counting Applications. Springer. Berlin. ISBN: 978-3-319-14929-5
Zhang, Z., Liu, Z. and Tian, Y. (2020). A DNA-Based FLIM Reporter for Simultaneous Quantification of Lysosomal pH and Ca2+ during Autophagy Regulation. iScience. 23(7): 101344.
Patting, M., Reisch, P., Sackrow, M., Dowler, R., Koenig, M. and Wahl, M. (2018). Fluorescence decay data analysis correcting for detector pulse pile-up at very high count rates. Opt Eng. 57(3): 031305.
Wahl, M., Röhlicke, T., Kulisch, S., Rohilla, S., Krämer, B. and Hocke, A. C. (2020). Photon arrival time tagging with many channels, sub-nanosecond deadtime, very high throughput, and fiber optic remote synchronization. Rev Sci Instrum. 91(1): 013108.
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Naumann, G., Lippmann, K. and Eilers, J. (2018). Photophysical properties of Na+‐indicator dyes suitable for quantitative two‐photon fluorescence‐lifetime measurements. J Microsc. 272(2): 136–144.
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Article Information
Publication history
Received: Sep 13, 2024
Accepted: Dec 3, 2024
Available online: Dec 17, 2024
Published: Feb 5, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Neuroscience > Neuroanatomy and circuitry > Fluorescence imaging
Neuroscience > Nervous system disorders > Cellular mechanisms
Cell Biology > Cell imaging > Live-cell imaging
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Determination of Dissociation Constants for the Interaction of Myosin-5a with its Cargo Protein Using Microscale Thermophoresis (MST)
RZ Rui Zhou
JP Jiabin Pan
XL Xiang-Dong Li
Published: Vol 15, Iss 3, Feb 5, 2025
DOI: 10.21769/BioProtoc.5176 Views: 49
Reviewed by: Chiara AmbrogioOm Prakash Narayan Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in eLIFE Jun 2024
Abstract
Myosin-5a (Myo5a) is an actin-dependent molecular motor that recognizes a diverse range of cargo proteins through its tail domain, playing a crucial role in the transport and localization of various organelles within the cell. We have identified a new interaction between Myo5a and its cargo protein melanophilin (Mlph), i.e., the interaction between the middle tail domain of Myo5a (Myo5a-MTD) and the actin-binding domain of Mlph (Mlph-ABD), by GST pulldown assay. We then intend to obtain the dissociation constant between Myo5a-MTD and Mlph-ABD using isothermal titration calorimetry (ITC) or microscale thermophoresis (MST), both of which are two commonly used methods for determining quantitative data on protein interactions. The advantages of MST over ITC include less protein usage, shorter operation time, and higher sensitivity. In this protocol, we present a method for using MST to determine the dissociation constants of Myo5a-MTD and Mlph-ABD, which were purified through overexpression in bacteria using affinity chromatography. The dissociation constant values obtained directly reflect the binding strength between these two proteins and provide a foundation for the isolation and purification of the complex in the future.
Key features
• A protocol for determining the dissociation constants between two purified proteins using microscale thermophoresis (MST).
• Detailed procedures for purification of recombinant proteins expressed in E. coli.
Keywords: MST Dissociation constant Protein interactions Protein expression Protein purification
Background
Myosin-5a (Myo5a) is an actin-dependent unconventional myosin that primarily functions as a transporter within cells. Among the various cargo proteins associated with Myo5a, melanophilin (Mlph) is currently the most extensively studied [1]. Mlph is a scaffold protein that interacts with small GTPase Rab27a at the N-terminal portion and Myo5a at the C-terminal portion, thus bridging Rab27a and Myo5a to form a trimeric complex.
Previous research has confirmed that Mlph contains two domains that bind directly to Myo5a: Mlph-GTBD, known as the globular tail domain (GTD)-binding domain, which interacts with the GTD of Myo5a; and Mlph-EFBD, known as the exon-F binding domain, which interacts with melanocyte-specific exon-F in the Myo5a tail [2]. The discovery of a third Myo5a-Mlph interaction enhances our understanding of the role that Myo5a plays in melanosome transport. In our initial study, we employed the traditional GST pulldown assay to investigate protein interactions, which qualitatively demonstrated the interaction between Myo5a-MTD and Mlph-ABD. However, considering that Mlph employs multiple structural domains to simultaneously interact with Myo5a [3–5], the GST pulldown assay is insufficient for comparing the binding affinities of the different structural domains of Mlph with Myo5a. In contrast, MST is a highly sensitive technique that can determine dissociation constants for ligand proteins by measuring fluorescence changes in the target protein [6].
MST is a technique used to determine dissociation constants for ligand proteins by quantifying the thermophoretic movement of fluorescent molecules in response to a temperature gradient. This method typically requires only nanomolar concentrations of target proteins, allowing for high sensitivity in detecting interactions. Additionally, the determination of MST is not influenced by the buffer composition in the system [7]. Many MST protocols utilize the overexpression of eGFP proteins in the cells as target proteins [8]. While this approach is simple and efficient, it may impact the conformation of the target protein. In our biological protocol, we performed affinity assays using MST by fluorescently labeling the amino groups of GST-tagged Mlph-ABD, which was expressed in E. coli and purified in vitro, as the target protein. The His-tagged Myo5a-MTD, also expressed in E. coli and purified in vitro, served as the ligand protein. This strategy allows us to arbitrarily select two proteins as the target and ligand proteins and to freely adjust their concentration, facilitating the success of the experiment. However, our method has limitations. For instance, the proteins used in the experiment must be easily purifiable to avoid loss of fluorescent labeling. Additionally, the concentration of fluorescent dyes may become excessive after labeling, as proteins typically contain multiple amino groups.
Materials and reagents
Reagents
1. IPTG (isopropyl β-D-1-thiogalactopyranoside) (Inalco, catalog number: 1758-1400)
2. Glutathione (GE Healthcare, catalog number: 17-5132-02)
3. Kanamycin (Inalco, catalog number: 1758-9316)
4. Ampicillin (Inalco, catalog number: 1758-9314)
5. Lysozyme (Solarbio, catalog number: L8120)
6. DNase I (Beijing DingGuoChangSheng, catalog number: DH113-5)
7. Glutathione-Sepharose 4 Fast Flow (GE Healthcare, catalog number: 17513201)
8. Ni-nitrilotriacetic acid agarose (QIAGEN, catalog number: 30210)
9. ATTO-488 NHS ester (ATTO-TEC GmbH, catalog number: AD488-35)
10. Coomassie brilliant blue G250 (Amresco, catalog number: M140-50G)
11. Tryptone (QiSong biology, catalog number: BQS133120)
12. Yeast extract (Oxiod, catalog number: LP0021)
13. Glucose (QiSong biology, catalog number: BQS119483)
14. Tris-base (Sigma, catalog number: T1387)
15. EDTA (Amresco, catalog number: 0105)
16. DTT (Amresco, catalog number: 0281)
17. β-Mercaptoethanol (Macklin, catalog number: M6230)
18. Imidazole (Sigma-Aldrich, catalog number: 0664)
19. Ethanol (Beijing Yili Fine Chemicals Co., Ltd)
20. Tween-20 (DingGuoChangSheng, catalog number: DH358-3)
21. DMSO (Leagene, catalog number: CC0118)
22. NaCl (Beijing Yili Fine Chemicals Co., Ltd)
23. MgCl2 (Beijing Yili Fine Chemicals Co., Ltd)
24. KCl (Beijing Yili Fine Chemicals Co., Ltd)
25. NaHCO3 (Beijing Yili Fine Chemicals Co., Ltd)
26. H3PO4 (Beijing Yili Fine Chemicals Co., Ltd)
Solutions
1. SOC (used for E. coli transformation) (see Recipes)
2. LB (used for E. coli culture) (see Recipes)
3. 10× TBS (used for equilibration of Ni-nitrilotriacetic acid-agarose or Glutathione-Sepharose 4 Fast Flow) (see Recipes)
4. GST-tag protein lysis buffer (used for GST-tagged protein purification) (see Recipes)
5. GST-tag protein washing buffer (used for GST-tagged protein purification) (see Recipes)
6. GST-tag protein elution buffer (used for GST-tagged protein purification) (see Recipes)
7. GST-tag protein dialysis buffer (used for GST-tagged protein purification) (see Recipes)
8. His-tag protein lysis buffer (used for His-tagged protein purification) (see Recipes)
9. His-tag protein washing buffer (used for His-tagged protein purification) (see Recipes)
10. His-tag protein elution buffer (used for His-tagged protein purification) (see Recipes)
11. His-tag protein dialysis buffer (used for His-tagged protein purification) (see Recipes)
12. Bradford (used for protein detection) (see Recipes)
13. MST buffer (used for dilution of fluorescently labeled proteins in the MST reaction) (see Recipes)
14. ATTO-488 NHS-ester dye (see Recipes)
Recipes
1. SOC
Reagent Final concentration Quantity or Volume
Tryptone 2% 20 g
Yeast extract 0.5% 5 g
NaCl 0.05% 0.5 g
KCl 2.5 mM 0.83 mL
MgCl2 10 mM 10 mL
Glucose 20 mM 20 mL
Adjust the final volume to 1 L with distilled water.
2. LB
Reagent Final concentration Quantity or Volume
Yeast extract 5 g/L 5 g
Tryptone 10 g/L 10 g
NaCl 10 g/L 10 g
Adjust the final volume to 1 L with distilled water.
3. 10× TBS
Reagent Final concentration Quantity or Volume
Tris-base 0.5 M 60.57 g
NaCl 1.5 M 87.66 g
Adjust the final volume to 1 L with distilled water. Adjust pH to 7.5 with concentrated HCl.
4. GST-tag protein lysis buffer
Reagent Final concentration Volume
Tris-HCl, pH 7.5 50 mM 4.5 mL
EDTA 2 mM 90 μL
DTT 1 mM 360 μL
Lysozyme 10 mg
Adjust the final volume to 90 mL with distilled water.
5. GST-tag protein washing buffer
Reagent Final concentration Volume
TBS 1× 50 mL
Adjust the final volume to 500 mL with distilled water.
6. GST-tag protein elution buffer
Reagent Final concentration Quantity or Volume
Tris-HCl, pH 8.0 50 mM 1 mL
NaCl 200 mM 1 mL
DTT 1 mM 20 μL
Glutathione 10 mM 0.06 g
Adjust the final volume to 20 mL with distilled water.
7. GST-tag protein dialysis buffer
Reagent Final concentration Quantity or Volume
NaHCO3 130 mM 10.92 g
NaCl 50 mM 2.92 g
Adjust the final volume to 1 L with distilled water. Adjust pH to 8.2–8.3 with concentrated HCl.
8. His-tag protein lysis buffer
Reagent Final concentration Volume
Tris-HCl, pH 7.5 50 mM 4.5 mL
EDTA 2 mM 90 μL
β-Mercaptoethanol 5 mM 30 μL
Lysozyme 10 mg
Adjust the final volume to 90 mL with distilled water.
9. His-tag protein washing buffer
Reagent Final concentration Volume
Tris-HCl, pH 7.5 50 mM 20 mL
Imidazole, pH 7.5 15 mM 6 mL
NaCl 150 mM 15 mL
β-Mercaptoethanol 5 mM 140 μL
Adjust the final volume to 500 mL with distilled water.
10. His-tag protein elution buffer
Reagent Final concentration Volume
Tris-HCl, pH 7.5 50 mM 1 mL
Imidazole, pH 7.5 250 mM 4 mL
NaCl 150 mM 750 μL
β-Mercaptoethanol 5 mM 5.6 μL
Adjust the final volume to 20 mL with distilled water.
11. His-tag protein dialysis buffer
Reagent Final concentration Quantity or Volume
Tris-HCl, pH 7.5 10 mM 10 mL
NaCl 200 mM 50 mL
DTT 1 mM 1 mL
Adjust the final volume to 1 L with distilled water.
12. Bradford
Reagent Final concentration Quantity or Volume
Coomassie brilliant blue G250 1.41 mM 100 mg
H3PO4 10% v/v 100 mL
Ethanol 5% v/v 50 mL
Adjust the final volume to 1 L with distilled water.
13. MST buffer
Reagent Final concentration Volume
Tris-HCl, pH 7.8 50 mM 2.5 mL
NaCl 100 mM 1.25 mL
MgCl2 10 mM 0.5 mL
Tween-20 0.05% v/v 125 μL
Adjust the final volume to 50 mL with distilled water.
14. ATTO-488 NHS-ester dye
Dissolve 5 mg of ATTO-488 NHS-ester dye powder in 510 μL of DMSO. Aliquot into 20 μL per tube, lyophilize, and store at -80 °C. Each tube contains 0.196 mg of ATTO-488 NHS-ester.
Laboratory supplies
1. General-purpose dialysis bags 8000–14000 (for protein purification ) (ShangHai Yuye, catalog number: MD1425)
2. Gravity flow column B (for column chromatography) (NanoTemper Technologies GmbH, catalog number: L001)
3. 200 μL PCR tubes (for MST experiments) (Axygen, catalog number: PCR-02-C)
4. Monolith capillaries (for MST experiments) (NanoTemper Technologies GmbH, catalog number: MO-K002)
Equipment
1. Ultrasonic cell pulverizer (for cell lysis) (NingBo XinZhi Biotechnology Co., LTD, model: JY92-II)
2. Ultracentrifuge (for rapid sample processing and preservation) (Beckman Coulter, model: Optima XPN-80)
3. Benchtop centrifuge (for precipitating insoluble proteins) (Eppendorf, model: 5415R)
4. Nanodrop-1000 (for measuring nucleic acid and protein concentrations) (Thermo Fisher)
5. NanoTemper® Monolith NT.115 (for MST measurements) (NanoTemper Technologies GmbH)
Software and datasets
1. Nanodrop-1000 4.64.0.0 (for determination of protein concentration) (Thermo Fisher Scientific)
2. Monolith® NT.115 (for analyzing MST data) (NanoTemper)
3. KaleidaGraph 4.0 (for data analysis and graphing) (Synergy)
4. Illustrator CS6 (for drawing images) (Adobe)
5. Mo.Control software (for controlling MST experiments) (NanoTemper)
Procedure
A. Protein expression
Expression of GST-Mlph-ABD
1. Transformation:
a. Transfer 100 ng of plasmids (1 μL) GST-Mlph-ABD/pGEX4T2 into 100 μL of BL-21(DE3) E. coli competent cells.
b. Keep the mixture on ice for 5 min, heat-shock at 42 °C for 45 s, and then immediately place back on ice for 2 min.
c. Add 200 μL of SOC medium to each sample and incubate at 37 °C with shaking for 45 min.
d. Spread 100 μL of the above culture to an ampicillin-resistant agar plate (LB agar with ampicillin) and incubate the plate at 37 °C for 12–16 h.
2. Culture of E. coli:
a. When the colonies on the plate grow to the appropriate size, pick up a single colony to inoculate 4 mL of the ampicillin-resistant LB medium (at room temperature) and incubate at 37 °C with shaking at 200 rpm for 6 h.
b. Use the above culture (~4 mL) to inoculate 250 mL of the ampicillin-resistant LB medium (room temperature) and incubate at 37 °C with shaking at 200 rpm for 3–4 h until the OD600 reaches 0.8–1.
c. Add 50 μL of 1 M IPTG (final concentration 0.2 mM) to the above culture to induce protein expression and incubate with shaking at 200 rpm at 37 °C for 3 h or at 17 °C for 12 h (General note 1).
3. Collect E. coli:
a. Harvest the induced E. coli by centrifugation at 4,000 rpm (~3,000× g) for 10 min at room temperature.
b. Resuspend the E. coli pellets with 1× TBS and precipitate the E. coli by centrifugation again at 4,000 rpm (~3,000× g) for 10 min at room temperature. Discard the supernatant and use the pellets for purification directly or store them at -80 °C for later use.
Expression of His-tagged Myo5a tail (His-Myo5a-MTD and His-Myo5a-MTDΔG)
1. Transformation:
a. Transfer 100 ng of bacterial expression plasmids (1 μL) His-Myo5a-MTD/pET30a or His-Myo5a-MTDΔG/pET30a into 100 μL of BL-21(DE3) E. coli competent cells (Myo5a-MTD can interact with Mlph, whereas Myo5a-MTDΔG cannot).
b. Keep the mixture on ice for 5 min, heat-shock at 42 °C for 45 s, and then immediately place back on ice for 2 min.
c. Add 200 μL of SOC medium to each sample and incubate at 37 °C with shaking for 45 min.
d. Spread 100 μL of the mixture to a kanamycin-resistant agar plate (LB agar with kanamycin) and incubate the plate at 37 for 12–16 h.
2. Culture of E. coli:
a. When the colonies on the plate grow to the appropriate size, pick up a single colony to inoculate 4 mL of the kanamycin-resistant LB medium (at room temperature) and incubate at 37 °C with shaking at 200 rpm for 6 h.
b. Use the above culture (~4 mL) to inoculate 250 mL of the kanamycin-resistant LB medium (room temperature) and incubate at 37 °C with shaking at 200 rpm for 3–4 h until the OD600 reaches 0.8–1.
c. Add 50 μL of 1 M IPTG (final concentration 0.2 mM) to the above culture to induce protein expression and incubate with shaking at 200 rpm at 37 °C for 3 h or at 17 °C for 12 h (General note 1).
3. Collect E. coli:
a. Harvest the induced E. coli by centrifugation at 4,000 rpm (~3,000× g) for 10 min at room temperature.
b. Resuspend the E. coli pellets with 1× TBS and precipitate the E. coli again by centrifugation at 4,000 rpm (~3,000× g) for 10 min at room temperature. Save the pellets and store at -80 °C or use them immediately for purification.
B. Protein purification
The purification of proteins requires several sequential steps, as shown in Figure 1.
Figure 1. Flowchart for protein purification
For the purification of GST-Mlph-ABD
1. Lysis:
a. Thaw the E. coli. pellet of GST-Mlph-ABD, which was collected from 250 mL of culture and stored at -80 °C.
b. Resuspend the E. coli pellets in 25 mL of GST-tag protein lysis buffer by pipetting up and down repeatedly and then stand on ice for 20 min.
c. Add 104 U/mL DNase I to achieve a final concentration of 10 U/mL, 1 M MgCl2 to a final concentration of 3 mM, and 4 M NaCl to a final concentration of 0.2 M. Mix by inverting and leave on ice for 10 min.
2. Sonication: Set the power of the ultrasonic cell pulverizer to 200 W, with a cycle of 3 s on and 7 s off, repeating this process 80 times for complete lysis of E. coli (General note 2).
3. Centrifugation: Centrifuge the lysate at 20,000 rpm (~40,000× g) using an ultracentrifuge for 40 min at 4 °C and save the supernatant of the lysate in a 50 mL conical tube.
4. Binding to GSH-Sepharose 4 Fast Flow beads (GSH-Sepharose beads):
a. Equilibrate GSH-Sepharose 4 beads: Suspend 1 mL of GSH-Sepharose 4 beads in 10 mL of 1× TBS; then, let the beads settle down and discard the supernatant.
b. Transfer the GSH-Sepharose 4 beads to the 50 mL conical tube containing the supernatant of the lysate and rotate the conical tube at 4 °C for 2 h.
5. Wash:
a. Centrifuge the 50 mL conical tube at 2,000 rpm (~800× g) for 10 min at 4 °C using a multifunctional tabletop centrifuge.
b. Discard the supernatant, then suspend the GSH-Sepharose 4 beads with ~50 mL of GST-tag protein washing buffer and centrifuge again at 2,000 rpm (~800× g) for 10 min. Discard the supernatant.
c. Transfer the GSH-Sepharose 4 beads to a disposal chromatography column. Rinse the column with GST-tag protein wash buffer and detect protein in the eluate by mixing 5 μL of eluate with 45 μL of Bradford staining solution. The color of the Bradford staining solution changes from brown to blue when it reacts with protein. Stop rinsing the column when the eluate does not change the color of Bradford staining solution.
6. Elution: Elute the target proteins using GST-tag elution buffer by gravity flow. Detect the proteins in the eluate by mixing 5 μL of eluate with 45 μL of Bradford staining solution. The color of the Bradford staining solution changes from brown to blue when it reacts with protein. Combine the eluate fractions containing high-concentration proteins.
7. Dialysis: Cut an appropriate length of the dialysis tube based on the estimated volume of the eluted protein. Place the dialysis tube in deionized water and boil at 100 °C for 3 min. Transfer the eluted proteins into the dialysis tube and dialyze against 1 L of GST-tag protein dialysis buffer overnight at 4 °C. This step is essential for labeling GST-Mlph-ABD with ATTO-488 NHS-ester, as it removes free amines.
8. Concentration:
a. Transfer the dialyzed proteins into ultrafiltration tubes and centrifuge at 4,000 rpm (~3,000× g) at 4 °C using a multifunctional tabletop centrifuge until the appropriate concentration is reached.
b. Aliquot the concentrated protein into a small volume (20–200 μL), quickly freeze in liquid nitrogen, and store at -80 °C.
c. Measure the protein concentration using Nanodrop-1000 and detect protein purity by SDS-PAGE (Figure 2). The purified GST-Mlph-ABD can be used for subsequent fluorescent labeling.
Figure 2. SDS-PAGE of purified GST-Mlph-ABD, His-Myo5a-MTD, and His-Myo5a-MTDΔG
Purification of His-tagged Myo5a tail (His-Myo5a-MTD and His-Myo5a-MTDΔG)
1. Lysis:
a. Thaw E. coli pellets of His-Myo5a-MTD or His-Myo5a-MTDΔG, which were collected from 250 mL of culture and stored at -80 °C.
b. Resuspend the E. coli pellets in 25 mL of His-tag protein lysis buffer by pipetting up and down repeatedly and then stand on ice for 20 min.
c. Add 104 U/mL DNase I to achieve a final concentration of 10 U/mL, 1 M MgCl2 to a final concentration of 3 mM, and 4 M NaCl to a final concentration of 0.2 M. Mix by inverting and leave on ice for 10 min.
2. Sonication: Set the power of the ultrasonic cell pulverizer to 200 W, with a cycle of 3 s on and 7 s off, repeating this process 80 times to fully lyse the organisms (General note 2).
3. Centrifugation: Centrifuge the lysate at 20,000 rpm (~40,000× g) using an ultracentrifuge for 40 min at 4 °C and save the supernatant of the lysate in a 50 mL conical tube.
4. Binding to Ni-nitrilotriacetic acid-agarose (Ni-agarose):
a. Suspend 1 mL of Ni-agarose in 10 mL of 1× TBS, let the beads settle down, and discard the supernatant.
b. Transfer the GSH-Sepharose 4 beads to the 50 mL conical tube containing the supernatant of the lysate and spin the conical tube on a rotary shaker at 4 °C for 2 h.
5. Wash:
a. Centrifuge the conical tube at 2,000 rpm (~800× g) for 10 min at 4 °C using a multifunctional tabletop centrifuge.
b. Discard the supernatant, then suspend the Ni-agarose with ~50 mL of His-tag protein washing buffer and centrifuge again at 2,000 rpm (~800× g) for 10 min. Discard the supernatant.
c. Transfer the Ni-agarose to a disposal chromatography column. Rinse the column with His-tag protein washing buffer and detect protein in the eluate by mixing 5 μL of eluate with 45 μL of Bradford staining solution. The color of Bradford staining solution changes from brown to blue when it reacts with protein. Stop rinsing the column when the eluate does not change the color of the Bradford staining solution.
6. Elution: Elute the target proteins using His-tag elution buffer by gravity flow. Detect the proteins in the eluate by mixing 5 μL of eluate with 45 μL of Bradford staining solution. The color of Bradford staining solution changes from brown to blue when it reacts with protein. Combine the eluate fractions containing high-concentration proteins.
7. Dialysis: Estimate the volume of the eluted protein and cut an appropriate length of the dialysis tube. Place the dialysis tube in deionized water and boil at 100 °C for 3 min. Transfer the eluted proteins into the dialysis tube and dialyze overnight at 4 °C in 1 L of His-tag protein dialysis buffer.
8. Concentration:
a. Transfer the dialyzed proteins into ultrafiltration tubes and centrifuge at 4,000 rpm (~3,000× g) at 4 °C using a multifunctional tabletop centrifuge until the appropriate concentration is reached.
b. Aliquot the concentrated protein into small volumes (20–200 μL), quickly freeze in liquid nitrogen, and store at -80 °C.
c. Measure the protein concentration using a Nanodrop-1000 and detect the protein purity by SDS-PAGE (Figure 2). The purified His-Myo5a-MTD and His-Myo5a-MTDΔG are directly used for MST experiments.
C. Labeling of GST-Mlph-ABD with ATTO-488 NHS-ester
1. Preparation of fluorescent dye: Dissolve 0.196 mg of ATTO-488 NHS-ester with 20 μL of DMSO to make a 10 mM solution, and then dilute to 1 mM by mixing 2 μL of ATTO-488 NHS-ester with 18 μL of GST-tag protein dialysis buffer.
2. Labeling:
a. Prepare 90 μL of 40 μM GST-Mlph-ABD protein by diluting GST-Mlph-ABD protein stock with GST-tag protein dialysis buffer.
b. Add 10.8 μL of 1 mM ATTO-488 NHS-ester dye to 90 μL of 40 μM GST-Mlph-ABD protein, resulting in a 3:1 molar ratio of ATTO-488 NHS-ester dye to GST-Mlph-ABD. Mix well.
c. Incubate for 1 h at room temperature in the dark (General notes 3–6).
3. Purification of the labeled protein:
a. Equilibrate the gravity flow column B with 3 mL of MST buffer three times.
b. Add a maximum of 500 μL of labeling reaction to the center of column B. Let the sample enter the column completely (when using less than 500 μL, adjust the volume to 500 μL using MST buffer after the sample has entered the column) and discard the flowthrough.
c. Add 2 mL of MST buffer to the center of column B and collect in 200–250 μL fractions.
4. Measure ATTO-488-labeled GST-Mlph-ABD concentration in effluent fractions using Nanodrop-1000:
a. Add 3 μL of deionized water dropwise to the Nanodrop's detection probe. Open the software Nanodrop-1000 and select the module Proteins & Labels.
b. Select the excitation light Alexa Fluor 488 for the fluorescent dye ATTO-488 NHS ester, add 3 μL of GST-tag protein dialysis buffer to the probe of the Nanodrop-1000, and click Blank.
c. Add 3 μL of effluent fractions to the Nanodrop-1000 probe and click Measure. The two blue highlight boxes in Figure 3 show the concentration of the fluorescent dye and protein.
d. Save the fractions containing high concentrations of ATTO-488-labeled GST-Mlph-ABD with a dye/protein molar ratio below 1.5 for subsequent MST experiments.
Figure 3. Measurement of the labeling efficiency of ATTO-488-labeled GST-Mlph-ABD using Nanodrop-1000
D. MST measurement of the interaction between His-Myo5a-MTD and GST-Mlph-ABD
1. Pre-test:
a. Dilute ATTO-488-labeled GST-Mlph-ABD to 40 nm with MST buffer.
b. Mix 10 μL of 40 nm ATTO-488-labeled GST-Mlph-ABD with 10 μL of His-tag protein dialysis buffer in a PCR tube, pipetting up and down several times to ensure adequate mixing, and then aspirate the mixture using Monolith capillaries (General note 7).
c. Put the capillaries in the slots on the sample tray, not touching the optical measurement section of the capillary.
d. Place the sample tray in the instrument by pushing it into the instrument tray slot as far as possible. The Monolith NT.115 instrument scans the tray and automatically determines the position of the capillaries on the tray.
e. Start the Mo.Control software and select Start New Session.
f. Select the green excitation filter for the upcoming experiment (General note 8).
g. Select the mode Pretest before performing a binding experiment and perform a pre-test experiment to detect the fluorescence intensity of the labeled GST-Mlph-ABD (see red highlight 1 in Figure 4). The fluorescence intensity of the target protein needs to be between 200 and 1,000, with a value between 400 and 800 being optimal.
Figure 4. Initial setup of an experiment
2. Sample loading:
a. Take 16 clean PCR tubes, labeled 1–16, and arrange them in order on the PCR tube holder.
b. Add 10 μL of His-Myo5a-MTD dialysis buffer to tubes 2–16 and 20 μL of 40 μM His-Myo5a-MTD to tube 1.
c. Dispense 10 μL of His-Myo5a-MTD from tube 1 and transfer it into tube 2, mixing thoroughly. Then, using the same tip, transfer 10 μL from tube 2 into tube 3. Repeat this process until tube 16. After thoroughly mixing the sample in the final tube 16, remove 10 μL from that tube and discard it (General note 9).
d. Add 10 μL of ATTO-488-labeled GST-Mlph-ABD to each of the 16 tubes, mix well, and centrifuge the samples at 13,000× g for 5 min. Transfer the sample from the PCR tubes into 16 clean Monolith capillaries sequentially.
e. Put the capillaries in the slots on the sample tray (General note 10).
f. Place the sample tray in the instrument by pushing it into the instrument tray slot.
3. Binding affinity measurement:
a. Return to the homepage of the Mo. Control software and select Binding Affinity (see red highlight 2 in Figure 4).
b. Enter the Plan interface and set the following parameters: the reaction temperature of the MST experiment, the initial and final concentrations of the target and ligand proteins, the estimated Kd, the type of capillary, the excitation power and the MST power (see red highlight in Figure 5A).
c. Click Go to Instructions (see blue highlight in Figure 5A) to enter the Instruction interface, which allows you to dilute the target and ligand proteins to the appropriate concentrations according to the protocol provided.
d. Click Start Measurement to enter the Results interface (see blue highlight in Figure 5B) and wait for the system to automatically measure the MST traces and the dissociation constant Kd.
Figure 5. Procedure for measuring binding affinity experiments using Mo.Control software. (A) Plan interface for setting the temperature of the reaction, initial and final concentrations of the target and ligand proteins, excitation power, and MST power. (B) Instruction interface that provides a protocol on how to dilute the target and ligand proteins to the desired concentration for binding.
Data analysis
The MST data can be analyzed with Monolith® NT.115, which comes with the MST machine, or using third-party software. Here, we show how to analyze MST data using the KaleidaGraph 4.0 software.
1. Input data:
a. Open the KaleidaGraph software and select File in the first icon from the top row of the KaleidaGraph software (Figure 6).
b. Select New under the File menu to enter the MST experiment data separately into the Data 1 table. Dose values should be entered in column A. The mean values of response obtained from three repetitions of MST experiments for two different sets of interacting proteins are entered in columns B and D. Columns C and E correspond to the standard deviation of the mean values in B and D over three experiments, respectively.
Figure 6. Screenshot of the KaleidaGraph version 4.0 software showing the location of the Data window under the File tab
2. Analysis data:
a. Select Gallery in the third icon from the top row of the KaleidaGraph software (Figure 7).
Figure 7. Screenshot of the KaleidaGraph version 4.0 software showing the submenu Scatter under the Linear menu under the Gallery tab
b. Select Scatter under the Linear menu to generate a dot plot.
c. Select Curve Fit in the sixth icon from the top row of the KaleidaGraph software and choose fit1 under the General menu (Figure 8).
Figure 8. Screenshot of the KaleidaGraph version 4.0 software showing the submenu fit1 under the General menu under the Curve Fit tab
d. Select the data in column B in the pop-up Curve Fitting Selection window. Click Define in the upper-right corner and enter the MST curve fitting formula (Figure 9):
m1 - m2/0.02/2*((m0 + 0.02 + m3) - ((m0 + 0.02 + m3) ^ 2 - 4*m0*0.02) ^ 0.5)/2; m1 = 953; m2 = 8; m3 = 0.5.
Figure 9. Screenshot of KaleidaGraph version 4.0 software showing the Curve Fitting Selection sub-window under the Curve Fitting Selection window
Assign the value of m1 as the maximum MST value, m2 as the maximum MST change, and m3 as the estimated Kd value in μM, which is the x-value corresponding to the vertical coordinate reaching half of the maximum of MST change (Figure 10) (General note 11).
Figure 10. Analysis of MST data using KaleidaGraph. Top, MST data plots and curve fits using KaleidaGraph. Middle, curve fitting equation and interpretation of related parameters. Bottom, the final figure used in publication (Pan et al. [9], https://elifesciences.org/articles/93662).
e. Click File and then save Graph as to save it in TIF format for editing in Illustrator CS6.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article:
• Pan et al. [9]. Identification of a third myosin-5a-melanophilin interaction that mediates the association of myosin-5a with melanosomes. eLife (Figure 3A).
We monitored the interaction between His-Myo5a-MTD and GST-Mlph-ABD using MST and obtained the dissociation constant (Kd) of 562 ± 169 nM of Myo5a-MTD for binding to Mlph-ABD. Consistent with the GST pulldown assay, we found that deletion of the C-terminal half of exon-G (His-Myo5a-MTDΔG) greatly decreased MST signaling.
General notes and troubleshooting
1. The optimal conditions for IPTG to induce the expression of different proteins vary, and factors such as the concentration of IPTG, E. coli culture temperature, E. coli culture time, and stain of E. coli need to be taken into account. In general, a lower IPTG concentration can reduce the burden of protein expression on host cells, while a higher IPTG concentration can induce protein expression more rapidly. Low-temperature induction can prolong the time of protein expression so that the protein can be folded sufficiently, but it may also cause protein degradation, while the opposite is true for high-temperature induction. In practice, the optimal conditions should be determined based on experimental requirements to achieve the best protein expression results. We routinely use 0.2 mM IPTG to induce protein expression.
2. After sonication, it can be observed that the bacterial solution becomes transparent; if it is still turbid, it may be that the lysis is not sufficient, and the volume of lysis buffer or the number of sonication needs to be increased.
3. Avoid using buffers containing primary amines (e.g., ammonium ions, tris, glycine, ethanolamine, glutathione) or imidazole for labeling fluorescent proteins, which compete with the labeled proteins to reduce labeling efficiency.
4. Concerning reducing agents, DTT and β-mercaptoethanol interfere with the labeling reaction and, therefore, need to be avoided. If a reducing agent is required during the labeling reaction, use TCEP. However, DTT and β-mercaptoethanol are better suited than TCEP for the subsequent MST experiment, since TCEP may in some cases reduce reproducibility.
5. Purified proteins containing carriers like BSA will not be labeled properly and should not be used.
6. Labeled protein concentration: If the protein concentration is too low, the labeling efficiency will be greatly affected, and the general concentration will not be lower than 0.5 mg/mL.
7. Place the capillary horizontally into the reaction tube to aspirate the sample. Do not touch the capillary in the middle section where the optical measurement will be performed.
8. The excitation color can be changed at the start of each new experiment.
9. Thorough mixing ensures consistent fluorescence intensity of fluorescent proteins in each PCR tube.
10. Note the order of the capillaries. The highest concentration is placed in the front of the tray. This position is denoted as “1” on the sample tray and in the control software.
11. To fit the equations correctly, it is necessary to make approximately correct guesses about m1, m2, and m3.
Acknowledgments
We are grateful to the core facility platform of the State Key Laboratory of Integrated Management of Pest Insects and Rodents at the Institute of Zoology for providing the laboratory equipment. This work was supported by the National Natural Science Foundation of China (31970657).
This protocol was adapted and modified from Pan et al. [4].
Competing interests
The authors declare that there are no competing financial interests.
References
Li, X. D., Ikebe, R. and Ikebe, M. (2005). Activation of myosin Va function by melanophilin, a specific docking partner of myosin Va. J Biol Chem. 280(18): 17815–17822.
Yao, L. L., Cao, Q. J., Zhang, H. M., Zhang, J., Cao, Y. and Li, X. D. (2015). Melanophilin Stimulates Myosin-5a Motor Function by Allosterically Inhibiting the Interaction between the Head and Tail of Myosin-5a. Sci Rep. 510874.
Fukuda, M. and Itoh, T. (2004). Slac2-a/melanophilin contains multiple PEST-like sequences that are highly sensitiveto proteolysis. J Biol Chem. 279(21): 22314–22321.
Geething, N. C. and Spudich, J. A. (2007). Identification of a minimal myosin Va binding site within an intrinsically unstructured domain of melanophilin. J Biol Chem. 282(29): 21518–21528.
Wu, X., Wang, F., Rao, K., Sellers, J. R. and Hammer, J. A. (2002). Rab27a is an essential component of melanosome receptor for myosin Va. Mol Biol Cell. 13(5): 1735–1749.
Nowak, P. M. and Woźniakiewicz, M. (2022). The Acid-Base/Deprotonation Equilibrium Can Be Studied with a MicroScale Thermophoresis(MST). Molecules. 27(3): 685.
Seidel, S. A., Dijkman, P. M., Lea, W. A., van den Bogaart, G., Jerabek-Willemsen, M., Lazic, A. and Duhr, S. (2013). Microscale thermophoresis quantifies biomolecular interactions under previously challenging conditions. Methods. 59(3): 301–315.
Magnez, R., Thiroux, B., Taront, S., Segaoula, Z., Quesnel, B. and Thuru, X. (2017). PD-1/PD-L1 binding studies using microscale thermophoresis. Sci Rep. 7(1): 17623.
Pan, J., Zhou, R., Yao, L. L., Zhang, J., Zhang, N., Cao, Q. J., Sun, S. and Li, X. D. (2024). Identification of a third myosin-5a-melanophilin interaction that mediates the association of myosin-5a with melanosomes. eLife. 13e93662.
Article Information
Publication history
Received: Oct 17, 2024
Accepted: Dec 3, 2024
Available online: Dec 17, 2024
Published: Feb 5, 2025
Copyright
© 2025 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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Confocal Live Imaging of Reproductive Organs Development in Arabidopsis
BW Binghan Wang
AB Amélie Bauer
AG Andrea Gómez-Felipe
SS Sylvia R. Silveira
DK Daniel Kierzkowski
Published: Vol 15, Iss 3, Feb 5, 2025
DOI: 10.21769/BioProtoc.5177 Views: 299
Reviewed by: Pooja VermaHeng ChenTasleem Javaid
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Cited by
Original Research Article:
The authors used this protocol in Plant Physiology Aug 2021
Abstract
Understanding how multicellular organisms are shaped requires high-resolution, quantitative data to unravel how biological structures grow and develop over time. In recent years, confocal live imaging has become an essential tool providing insights into developmental dynamics at cellular resolution in plant organs such as leaves or meristems. In the context of flowers, growth tracking has primarily been limited to sepals, the outermost floral organs, or the post-fertilization gynoecium, which are easily accessible for microscopy. Here, we describe a detailed pipeline for the preparation, dissection, and confocal imaging of the development of internal reproductive floral organs of Arabidopsis thaliana including both the stamen and gynoecium. We also discuss how to acquire high-quality images suitable for efficient 2D and 3D segmentation that allow the quantification of cellular dynamics underlying their development.
Key features
• Fine dissection of tiny and tightly enclosed floral organs.
• Confocal live imaging method allowing long-term observation of plant reproductive morphogenesis.
• Assessing the quality of acquired images for efficient segmentation at cellular resolution in 2D and 3D.
Keywords: Confocal microscopy Live imaging Floral organs Stamen Gynoecium Organogenesis
Background
Flowers are essential for plant reproduction and vital for crop yield. The mature flower of Arabidopsis thaliana consists of concentric whorls of floral organs: non-reproductive sepals and petals that surround stamens and carpels (fused together to form gynoecium). The stamen, the male reproductive organ of the flower, consists of anther-producing pollen that is supported by a long filament. The gynoecium, the female reproductive structure, is composed of two fused carpels (forming the ovary) topped by the style and pollen-receiving stigma. After fertilization, the gynoecium develops into fruit, which is crucial for plant propagation.
The development of floral reproductive organs is difficult to observe, as early emerging sepals quickly enclose the initiating stamens and gynoecium making them inaccessible for imaging without dissection. Widely used methods relying on tissue fixing, sectioning, or clonal analysis have provided valuable insights into the development of internal floral organs [e.g., 1–5]. However, these methods lack the spatial and temporal resolution to understand and uncover dynamic developmental processes underlying organogenesis at cellular resolution. While confocal live imaging has been extensively used to follow growth in easily accessible sepals [e.g., 6–10], such an approach has only recently been applied to uncover developmental dynamics in reproductive organs [e.g., 11–13].
In this paper, we describe the confocal live imaging method to observe the growth of Arabidopsis thaliana stamen and gynoecium. We detail the procedures for dissecting and exposing internal floral organs and acquiring confocal images of the same sample over several consecutive days. Furthermore, we provide some hints on how to increase the survival rate of the sample and improve the quality of confocal images suitable for 2D or 3D segmentation. Our protocol enables observations with minimal perturbation of the normal in planta morphogenesis allowing long-term quantitative observation of these hidden floral organs at cellular resolution.
Materials and reagents
Biological materials
1. Reproductive organs of 4-week-old Arabidopsis thaliana Col-0 carrying a plasma membrane–localized fluorescent marker. Here, we used plants carrying pUBQ10::myr-YFP construct [14]; however, different plasma membrane marker lines can be used
Reagents
1. Murashige and Skoog basal salt mixture (Sigma, catalog number: M5524-50L)
2. Sucrose (Fisher, catalog number: S5-3)
3. Murashige and Skoog vitamin solution (Sigma, catalog number: M3900-50ML)
4. Agar (Fisher, catalog number: BP1423-2)
5. Plant preservative mixture (PPM) (Plant Cell Technology, catalog number: 71806-1)
6. 95% denatured alcohol (Fisher, catalog number: HC-1100-1GL)
7. Propidium iodide (PI) (Sigma, catalog number: P4170) (potential carcinogen)
Solutions
1. 1/2 MS medium (for 1 L) (see Recipes)
2. 0.1% PPM solution (see Recipes)
3. 70% ethanol solution (see Recipes)
4. 0.1% PI staining solution (see Recipes)
Recipes
1. 1/2 MS medium (for 1 L)
2.15 g of Murashige and Skoog basal salt mixture
10 g of sucrose
1 mL of Murashige and Skoog vitamin solution
Add deionized H2O to 1 L
Adjust pH to 5.8
15 g of agar
2. 0.1% PPM solution
1 L of sterile deionized H2O
1 mL of plant preservative mixture (PPM)
3. 70% ethanol solution
737 mL of 95% denatured alcohol
263 mL of sterile deionized H2O
4. 0.1% PI staining solution (potential carcinogen)
1 mg of propidium iodide
1 mL of deionized H2O
Laboratory supplies
1. 35 × 10 mm Petri dishes (SARSTEDT, catalog number: 82.1135.500)
2. Laboratory film (ParafilmTM)
3. 200 μL and 1 mL pipette and corresponding tips
4. Precision tweezers with fine point (Dumont No. 5)
5. Syringe needles, 25 G × 1" (BD®, catalog number: 305125)
6. Tungsten 1 µm probe tips (Lambda, catalog number: T20-10)
7. Low lint tissue wipe (KimwipesTM, Kimberly-Clark)
8. Deionized H2O
9. Scalpel blades (FEATHER, # 15)
10. Surgical tape (MicroporeTM tape 3M)
11. Plastic pots, trays, and lids for plant potting (thermoformed square pots with drainage 2.63" × 2.63" × 2.25", no-hole trays 21" × 11" × 2.5", and compatible plastic dome 21.50" × 11" × 2.10")
12. All-purpose growing soil mixture (ASB Greenworld Grower Mix)
Equipment
1. Autoclave
2. Laminar flow cabinet
3. Dissecting stereomicroscope with a minimum of 4× zoom magnification (Zeiss, model: Stemi 35)
4. Upright confocal microscope (Zeiss, model: LSM800) equipped with long working distance, water-dipping lenses with a good numerical aperture (W Plan-Apochromat 40×/1.0 DIC M27 FWD = 2.5 mm)
5. Growth chamber (Conviron GEN1000)
Software and datasets
1. Zeiss Zen 2.6 blue edition (Carl Zeiss Microscopy GmbH)
Procedure
A. Plant growth
1. Sow the seeds in the pots with wet soil. Keep a distance of at least 5 cm between seeds. We suggest sowing five seeds per pot, one corner each and one at the center.
2. Accommodate the pots in trays with a 1 cm layer of water and cover with the lids to keep humidity.
3. Keep the trays with lids at 4 °C for 48 h of vernalization.
4. Transfer covered trays to the growth chamber set to long-day conditions (16 h of illumination, around 150 μmol m-2·s-1), with 60%–70% relative humidity at 22 ± 1 °C.
5. Remove the plastic lids after around 7 days, when the first two leaves are visible to the naked eye.
6. Water trays every other day by pouring around 1 cm of water into the bottom of the tray.
Note: Plants usually start bolting after around 3 weeks and are ready for dissection after around 4 weeks.
B. Preparation of imaging and culture plates
1. Prepare 1/2 MS medium (see Recipes).
2. Autoclave for 30 min at 120 °C.
3. Let the medium cool to around 50 °C before adding 0.1% PPM solution (see Recipes).
Note: PPM contains preservative and biocide agents that inhibit the germination of bacteria and fungal spores in in vitro plant cultures.
4. Under the laminar flow cabinet, fill 1/2–2/3 of the 35 × 10 mm Petri dishes with medium and wait for it to solidify.
5. Wrap the plates with parafilm to avoid contamination and drying.
6. Store plates at 4 °C for up to three months.
C. Preparation for dissection
1. Bring the Petri dishes containing the 1/2 MS medium to room temperature to avoid cold shock when placing the dissected sample.
2. Clean working surfaces, tweezers, scalpels, and dissection needles with 70% EtOH solution. It is not necessary to work in a sterile environment like a laminar flow cabinet but ensure your hands, area, and tools are always clean.
3. Place a Kimwipe moistened with deionized H2O on the dissection stereoscope stage.
Notes:
a. Other types of tissue wipes may be too soft and may easily tear apart when wet during dissection.
b. Be mindful of the amount of deionized H2O applied to the tissue. Too much water will soak the sample and make dissection difficult; if not enough water is applied, the sample will dehydrate (Video 1). Re-apply water whenever the Kimwipe is dry.
Video 1. Moistening the tissue wipe. How to prepare for dissection, applying the correct amount of water to the tissue wipe.
D. Dissection
Note: Ensure plants are well-watered, especially one day before the dissection.
1. Choose 4-week-old plants (Figure 1A). Pick an inflorescence that has already formed around 10 mature siliques.
Notes:
a. This is a desirable stage because the inflorescence presents enough young flower buds to be selected for dissection while not being surrounded by too many older flower buds. Additionally, at this stage, you have a stem that is long enough to hold and reposition the sample during dissection.
b. Older plants will be less vigorous and not suitable for imaging.
Figure 1. Preparation for live imaging. A–G. Dissection of floral organs. Inflorescence before (A) and after (B) removal of older flower buds. C. Flower bud at floral stage 8. The red dashed line indicates the base of the sepal. D. Schema of flower bud removal using a needle. The red dashed line indicates the location of the cutting. The curved arrow represents the direction of the movement done with the needle to remove the sepal. E. Dissected flower bud after removal of medial and lateral sepals. F. Dissected flower bud after removal of sepals and two long stamens exposing the gynoecium. G. Earlier dissected flower buds after medial sepal removal exposing stamen primordia. Dotted lines indicate the silhouette of developing organs, stamens (red) in (E) and (G), and gynoecium (blue) in (E) and (F). H. Placement of samples in a pre-cut chamber in the in vitro medium. Scale bars = 1 cm in (A), 200 μm in B, 100 μm in (C, E–G).
2. Prior to dissection, clean hands with 70% EtOH solution.
3. Cut the inflorescence from the main branch with a clean razor blade or scissors. Make sure you leave around 2–3 cm of the stem so you can hold it with one hand while dissecting with the other.
4. Place the stem on the wet Kimwipe. Grip the stem with the index finger of your non-dominant hand, pressing lightly so as not to crush the sample. Hold the dissection tool (tweezer or needle) with your dominant hand.
Note: During dissection, you should be able to gently roll the stem by slowly moving your finger left and right without crushing the stem against the tissue (Videos 2 and 3).
Video 2. Removal of older flowers. How to remove older flower buds with tweezers while rolling the inflorescence stem.
Video 3. Removal of younger flowers. How to remove younger flower buds with needles while rolling the inflorescence stem.
5. Remove older flowers around the spiral, rolling the inflorescence until you reach the desired stage (Figure 1B and Video 2). The oldest and more distant flowers can be removed by breaking the pedicel with tweezers (Video 2) or cutting with a razor or scalpel blade. Younger flower buds that are closer to the bud of interest should be cut with needles (Video 3).
Notes:
a. The remaining pedicel segments from removed flower buds can later be used to reposition samples avoiding directly touching the stem. However, if the segments are too many and too long, they may hamper the rolling of the stem.
b. If you are uncertain what flower bud size corresponds to which floral stage, carefully open the flower buds from outer to inner (bigger to smaller) until you reach the desired stage. A staging reference can be found in Smyth et al. [1], and cellular resolution sizes and landmarks can be found in references [11] and [12].
6. Roll the stem with your finger to position the flower of the desired stage facing upward.
7. With a dissection needle, scratch or poke the base of the medial sepal as close as possible to the junction between the sepal and pedicel to facilitate sepal detachment (Figure 1C, D and Video 4).
8. Carefully remove the medial sepal without touching other tissues. We suggest using the tip of the needle to lift up the distal part of the sepal and fold it toward the base to break at the base (Figure 1D and Video 4). One or two lateral sepals can be removed in the same way (Figure 1E and Video 4). Alternatively, cut the sepal directly from the base by pushing it up with the tip of the needle (Video 4).
a. If imaging stamens (Figure 1E, F), be careful not to poke the sepal too deep and damage the base of the stamen.
b. If imaging the gynoecium, also remove the two long stamens (Figure 1G and Video 4).
Notes:
a. To facilitate sepal removal, you can gently scratch the tip of the needle onto a hard surface to create a hook. The hook can help pull the tip of the sepals.
b. Depending on the floral stage, the two medial petal primordia and lateral stamens may be present but may not yet cover the organs of interest. Keeping them may increase the survival rate of the dissected bud at very early stages. These organs can be removed with tweezers or a needle at any point during the experiment when they extend and partially cover the organ of interest.
c. If the organ primordia that you want to remove are too small to be pulled with the tweezers or cut without damaging the tissue behind, or if there is any organ residue attached to the bud, use the dissection needle to poke and damage it. This will impair their growth, preventing them from hiding the organs of interest later in the time-lapse.
Video 4. Flower bud dissection. Tricks to remove sepals and stamens that enclose the organ of interest.
9. Gently remove most of the remaining younger flower buds by chopping them off with needles. Only keep a few buds behind the dissected one as physical support (Video 4). This will increase the survival rate of dissected buds. The supporting flower buds can be removed later once the development of the targeted organ stabilizes.
10. Cut the stem around 1 cm from the tip (Video 5).
Notes:
a. To ensure sample health and survival, make sure to have a sharp and clean cut removing the stem extremity that may have been pressed by the finger or hurt by tweezers.
b. The stem grows during long-term live imaging. A longer stem brings extra difficulties for positioning. However, cutting the stem afterward may impact the plant’s survival. To avoid having to cut the stem as it grows, try to keep the stem shorter from the start.
Video 5. Final chop. How to make the final cut on the stem before mounting the sample.
11. Mount the stem into the MS plate.
a. If imaging the dissected flower bud horizontally, use a scalpel to cut a chamber in the medium and a channel perpendicular to the chamber to place the dissected stem (Video 6). Position the sample perpendicular to the microscope lens (Video 7).
b. If imaging the dissected flower bud vertically, use a needle to make a hole in the agar and insert the sample by the stem. More about choosing sample mounting positions will be discussed in section F, step 4c, note b.
Notes:
a. When mounting and repositioning the sample, touch the pedicel residue and try to avoid touching the stem directly. If touching the stem is necessary, apply as little pressure as possible, rubbing the surface of the stem instead of pinching it.
b. Each Petri dish can initially accommodate up to around five inflorescences (Figure 1H). As the samples become big, we suggest reducing the sample number per Petri dish to avoid a longer time of submersion while the other samples on the same plate are being imaged.
Video 6. Preparation for sample mounting. How to cut a chamber and a channel in the agar to mount the sample for horizontal imaging.
Video 7. Mounting the sample. How to properly mount the sample on the imaging plate. Tricks to position and reposition the sample by rotating it.
E. Preparation for imaging
1. Fill in the cut-out chamber with PPM solution (see Recipes), ensuring all samples are completely submerged in the solution.
Note: If there is a bubble in front of the organ of interest, use a 200 μL pipette, ensure the tip is targeted on the dissected flower bud, and gently pipette a few times to remove the bubble.
2. Leave the sample and the medium submerged in the solution for a few minutes (5–10 min) to stabilize in the immersion solution prior to imaging. Plant tissue and agar may expand during water intake, leading to the displacement of the organ during a confocal scan.
F. Imaging
1. Place the Petri dish on the confocal stage and adjust the Petri dish holder (Figure 2A).
Figure 2. Imaging for growth quantification analysis. A. Setup of a Petri dish with PPM solution under the water immersion objective. B. Confocal image of dissected flower bud exposing gynoecium. Blue dotted lines indicate the same gynoecium indicated in Figure 1F. C. Confocal image of dissected flower bud exposing stamen primordia. Red dotted lines indicate the same stamen primordia indicated in Figure 1G. D. Medial confocal Z-section of dissected flower buds. Gynoecium imaged with oversaturated pixels in the outermost layers and stamen primordia imaged with oversaturated and unsaturated pixels in the inner layers. E. Comparison of Z-axis stack size between non-moving and an extreme case of moving sample. Red dotted lines indicate the same stamen primordia indicated in (C). F–G. Schema of samples mounted vertically (F) and horizontally (G) in the medium and corresponding resulting confocal images. White dashed lines indicate the position of the transversal digital cross-sections of the developing stamen. The arrow in (F) indicates the limitation of signal acquisition in depth. H. Confocal image of a developing gynoecium composed of stitched stacks (green and red). White square outlines the first stack, and the dashed square outlines the second stack. Note the overlapping area. I. Confocal image of stained developing gynoecium with split channels. Plasma membrane marker (PM) (top in green) and propidium iodide (PI) (bottom in red). Arrows indicate damage caused during dissection. Long colored arrows in (C) and (E–G) indicate the axes of the reconstructed confocal images X (blue), Y (yellow), and Z (pink). Scale bars = 20 μm in B, C, and E–G, 50 μm in D, H, and I.
2. Select the long working distance water immersion objective (e.g., W Plan-Apochromat 40×/1.0).
Note: In our system, we use W Plan-Apochromat 40×/1.0, but other water-dipping lenses and magnifications can be used as long as they have a high numerical aperture (we suggest NA > 0.8).
3. Set up the excitation and signal collection range according to the fluorophore in question (for the plasma membrane marker used here, pUBQ10::myr-YFP excitation was performed using a diode laser with 488 nm and the signal was collected at 500–600 nm).
4. After locating and focusing the sample, switch to live mode and adjust gain and laser power to visualize the contours of cells on the screen. Explore the sample in x, y, and z axes and set up image acquisition parameters as well as the Z-stack range. Acquire your confocal image of the organ of interest at the desired initial developmental stage (Figure 2B, C).
Critical: Start your scan from a Z-position above the organ surface (before cells become visible) to make sure the entire organ surface is imaged.
For cell segmentation and growth quantification with the software MorphoGraphX [16] or similar, image acquisition should be done with the following parameters:
a. Acquire 16-bit images. A larger color range helps to obtain more information from darker areas of the image.
b. Define a small Z-step size. For best results, adjust the Z-step size according to the x and y resolution, keeping a voxel size as close as possible to cubic.
Note: A step size equal to or smaller than 1 μm and 0.5 μm are recommended for 2D and 3D segmentation, respectively.
c. Optimize image contrast by adjusting laser power and master gain.
For cell segmentation, it is important to have a good contrast between the inside of the cell and cell outlines. In this case, saturating the plasma membrane signal is recommended. Adjust laser power and master gain while monitoring pixel saturation through the range indicator tool from the confocal imaging software (on the Zeiss LSM800 used here, power was around 2% and gain was 650 V). Make sure to make the same adjustments along the Z-stack, either by gradually increasing pixel saturation while scanning or, if your imaging software allows, setting different laser power and master gain values for different Z-positions, prior to starting acquisition.
Notes:
a. For surface (2.5D) segmentation, adjust the laser to oversaturate pixels at the outermost layer and ignore the signal inner layers even if it is weak (Figure 2D gynoecium). However, if inner tissues are also of interest for 3D analysis, adjustments must be made while paying attention to all cell layers (Figure 2D, stamen).
b. According to your research goal, you may have to optimize the positioning of the sample for a better signal of the region of interest (Figure 2F, G). Imaging the sample vertically ensures higher signal quality in inner tissues but impairs acquisition of the complete organ length (Figure 2F), while imaging horizontally ensures the acquisition of the entire front organ surface, with lower image quality in internal tissue (Figure 2G).
Caution: Be careful when increasing laser power. Exposure to high laser intensity may cause photobleaching and stresses that are harmful to plant growth.
c. Keep in mind that if your research goal involves quantifying fluorescence levels and comparing fluorescence between samples, the signal saturation should be avoided.
d. There is always a trade-off between image quality for segmentation, laser exposure tolerance, and the available time for the experiment. The experiment setup should be chosen based on the specific goals and work conditions.
5. After taking the confocal image, click on the 3D display tool from the imaging software to check if the stack acquired represents the realistic 3D shape of the organ.
Notes:
a. The samples may move during image acquisition for several reasons, e.g., strong vibrations, physical movement of the sample, and water absorption by the agar. These movements result in deformed 3D stacks (an example of an extreme case of flattening in the Z-axis is shown in Figure 2E). The data extracted from a deformed confocal stack will not be accurate and representative of reality.
b. To avoid movement, we recommend 1) always checking the functioning of the antivibration platform in which the microscope is installed before every imaging session, and 2) respecting the recommended time of incubation prior to imaging (see step 2 section E).
c. If strong movement is detected even after the incubation period, transfer the sample to a new Petri dish with a newly cut chamber and channel (see step 11 section D) and carefully position the stem on it.
6. For samples that are larger than the field of view, acquire multiple overlapping stacks.
Notes:
a. It is recommended that the acquisition be started from one distal corner of the organ. If the complete organ width is acquired in one stack, move the stage longitudinally (along the organ's proximal distal axis) to continue scanning the length of the organ. If the tissue is broader, keep moving and scanning horizontally (medial–lateral axis) until the complete organ width is acquired before moving longitudinally again. Make sure to have a small overlapping area between each stack to guide you in reconstructing the complete image.
b. If multiple confocal stacks are acquired for the same sample, use image analysis software of your preference to roughly stitch all the stacks immediately after scanning and confirm the entirety of the sample was imaged. If the final image has any missing pieces, it is still possible to redo the imaging (Figure 2H).
c. Depending on scanning speed, Z-stack depth, and Z-step size, each confocal stack may take 2–5 and 5–10 min to scan for 2D and 3D analysis, respectively. If a sample requires overlapping stacks, the acquisition time is multiplied by the total number of stacks (see General notes 5–6).
G. Post imaging
1. Unload the Petri dish from the microscope stage, being careful not to drop the PPM solution on the microscope.)
2. Discard the PPM solution. Remove the excess liquid from the agar chamber with a Kimwipe without touching the analyzed organ.
Critical: Keeping even a small volume of liquid around the sample will impair its growth.
3. Verify that the stem is in the correct position (not tilted/incorrect angle). Reposition it if necessary (Video 7).
4. Seal the Petri dish with Micropore tape.
5. Cultivate the dissected samples in an in vitro room/cabinet with the same controlled growth conditions as the growth chamber for the entire duration of the experiment. Place the Petri dish in a position that ensures negative gravitropism (vertical if the sample is horizontal, horizontal if the sample is vertical).
Note: If the experiment lasts for more than one week, at the end of the first week, place the sample into a new Petri dish with fresh medium to avoid contamination and to provide enough nutrients.
6. After the defined time interval (e.g., 24 h), restart from step E1.
Data analysis
In this protocol, we describe in detail a live-imaging method that enables tracking the development of Arabidopsis reproductive organs. Unlike previous live-imaging studies, which are generally limited to easily accessible floral organs namely sepals, our protocol allows the detailed imaging of the innermost reproductive floral organs for extended periods of time (up to two weeks). By carefully dissecting the external floral whorls, we expose the internal floral organs and observe their development from early initiation until the establishment of their final morphology, while preserving in vivo–like characteristics of the organogenesis (Figure 3).
Data obtained with this protocol can be used for both 2D [11,12] and 3D [13] cell segmentations and growth quantifications with advanced image analysis software such as MorphoGraphX [15,16]. Furthermore, this protocol may be reproduced to track the development in both wild-type and mutant specimens or samples upon chemical treatments, facilitating the understanding of how molecular factors modify key cellular behaviors during organogenesis. The method was successfully applied using Arabidopsis mutants with defects in critical reproductive processes, such as carpel fusion, style identity [12], and sporogenesis [13]. All these mutants survived the conditions imposed by this method, as did the wild type, while retaining their characteristic defective phenotypes. Beyond Arabidopsis, we believe that this protocol could be adapted for long-term, quantitative studies of reproductive organ development in other species, which are currently limited to either early primordia initiation [21] or gynoecium shape transformations post-fertilization [22]. Since the method depends on successful in vitro culture, optimization of the culture medium may be necessary, depending on the species' sugar and nutrient requirements. An additional challenge may arise with bigger organ sizes, as larger species may not fit within our recommended setup and will likely extend the duration of the experiments. Overall, this protocol offers a robust and adaptable approach, with the potential to significantly enhance our understanding of morphogenesis in the reproductive organ of the flower.
Figure 3. Time-lapse live imaging of A. thaliana internal floral organs. A. Confocal images of a time-lapse series of the developing stamen from 1 to 10 days after primordia initiation [11]. B. Confocal images of a time-lapse series of the developing gynoecium from 4 to 13 days after primordia initiation [12]. Scale bars = 100 μm.
Validation of protocol
This protocol or parts of it has been used and validated in the following research articles:
• Silveira et al. [11]. Live-imaging provides an atlas of cellular growth dynamics in the stamen. Plant Physiology.
• Gómez-Felipe et al. [12] Two orthogonal differentiation gradients locally coordinate fruit morphogenesis. Nature Communications.
• Kierzkowski et al. [13] Mechanical interactions between tissue layers underlie plant morphogenesis. Preprint available at Research Square.
General notes and troubleshooting
1. During several steps of dissection, while manipulating the sample with tweezers and needles, non-intentional damage to the plant tissue may occur. Samples may recover, but such wounds may impair your experiment (e.g., locally affect growth). Therefore, to increase the dissection success rate, it is essential to practice before directly investing time into the time-lapse live imaging. It usually takes one to several weeks before you are able to confidently dissect depending on the initial size of your samples.
2. During the practice period, propidium iodide (PI) staining is a useful tool to check for mistakes in dissection. Because PI penetrates wounded cells, it facilitates spotting the location of tissue damage, which is not always visible with the plasma membrane channel (Figure 2I). If choosing to use this strategy, we recommend staining samples for 5 min with 0.1% PI solution (see Recipes). Staining can be performed before mounting the sample in the medium by applying a few drops of PI solution directly to the dissected flower on a glass slide. Alternatively, after mounting the sample, fill the Petri dish or agar chamber with the 0.1% PI solution. In both cases, after 5 min, rinse it twice with PPM solution before filling the Petri dish for imaging. It is recommended that the acquisition be started from one distal corner of the organ. If the complete organ width is acquired in one stack, move the stage longitudinally (along the organ's proximal–distal axis) to continue scanning the length of the organ. If the tissue is broader, keep moving and scanning horizontally (medial–lateral axis) until the complete organ width is acquired before moving longitudinally again. Make sure to have a small overlapping area between each stack to guide you in reconstructing the complete image.
Caution: Propidium iodide is a potential carcinogen and should be handled with care. Avoid contact with skin and eyes by wearing suitable protective clothing, gloves, and eye/face protection. Dispose of the dye safely following local regulations.
3. In some cases, the dissected sample may exhibit limited growth between the first two time points. However, it may recover and develop normally later on. Therefore, it is recommended not to discard the sample immediately but to continue imaging and allow additional time for recovery. This approach can improve the sample survival rate and reduce the overall difficulty of experiments.
4. While this method has been shown to support all developmental stages (Figure 3), it is not entirely non-invasive. Stress from dissection, in vitro cultivation, and laser exposure can impact development. For example, exposing a primordium earlier may lead to earlier cessation of growth. To ensure a comprehensive and accurate time-lapse series, we recommend having as many replicates as possible and then selecting a minimum of three, based on the quality of the series, for the following analysis. When evaluating the quality of a time-lapse series, consider the following criteria: 1) Series that best represent the general developmental patterns expected based on literature and observed frequently across all replicates. 2) Series that capture the full or most of the developmental stages of interest within a single time-lapse. 3) Series with minimal to no cell damage at individual time points. If there are less than three, perform the experiment again following the same principle.
5. If fewer than three replicates are obtained, repeat the experiment to collect more. If needed, combining two independent partially overlapping time-lapse series can serve as a single replicate, offering a comprehensive view of organ development.
6. As samples grow bigger, the time to image one sample increases as days go by. Consider this when scheduling microscope slots and planning how many replicates will be followed at a time. For instance, the gynoecium at 13 DAI shown in Figure 3 is composed of approximately 25 confocal stacks. Therefore, imaging the whole organ at later developmental stages could take more than 1 h for a single sample.
7. Experimental setup decisions should be made by carefully weighing all relevant trade-offs, taking into account the specific conditions of your study and the requirements of your research questions. For instance, consider whether the research group has unrestricted or flexible access to a confocal microscope or if it is essential to capture the entire organ or the full developmental process.
8. The images obtained using this method were shown to be suitable for quantitative analysis in both 2D and 3D using the image analysis software MorphoGraphX [15–16]. The studies that validate this protocol outline some processes and parameters used for image processing, surface extraction, cell segmentation, and growth quantification in their method sections [11–13]. Detailed step-by-step instructions for analysis in MorphoGraphX can be found in several method and protocol publications [17–20]. Additional recommendations, information on the required computer hardware, tutorial videos, and troubleshooting resources are available at https://morphographx.org/. We also suggest visiting https://forum.image.sc/tags/MorphoGraphX and engaging in discussions.
Acknowledgments
We thank Viraj Alimchandani for critical reading of the manuscript.
This work was supported by Discovery grant (RGPIN-2018-05762) from the Natural Sciences and Engineering Research Council of Canada, as well as NOVA grant from the Fonds de Recherche du Québec Nature et Technologies and Natural Sciences and Engineering Research Council of Canada (2023-NOVA-327566).
This protocol was adapted and modified from Silveira et al. [11] and Gómez-Felipe et al. [12].
Competing interests
Authors declare that they have no competing interests.
References
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Article Information
Publication history
Received: Aug 29, 2024
Accepted: Nov 27, 2024
Available online: Dec 17, 2024
Published: Feb 5, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Plant Science > Plant developmental biology > Morphogenesis
Plant Science > Plant cell biology > Cell imaging
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The authors used this protocol in The Journal of Cell Biology Sep 2022
Abstract
Macrophages are known for engulfing and digesting pathogens and dead cells through a specialized form of endocytosis called phagocytosis. Unfortunately, many macrophage cell lines are refractory to most reagents used for transient transfections. Alternative transient approaches, such as electroporation or transduction with lentiviral vectors, typically cause cell death (electroporation) or can be time-consuming to generate numerous lentivirus when using different genes of interest. Therefore, we use the Sleeping Beauty system to generate stably transfected cells. The system uses a “resurrected” transposase gene named Sleeping Beauty found in salmonid fish. Experimentally, the system introduces two plasmids: one carrying the Sleeping Beauty transposase and the other with an integration cassette carrying the gene of interest, a reverse-doxycycline controlled repressor gene, and an antibiotic resistance gene. The construct used in this protocol provides puromycin resistance. Stable integrations are selected by culturing the cells in the presence of puromycin, and further enrichment can be obtained using fluorescence-activated cell sorting (FACS). In this protocol, we use the Sleeping Beauty transposon system to generate RAW264.7 cells with doxycycline-inducible inositol polyphosphate 4-phosphatase B containing a C-terminal CaaX motif (INPP4B-CaaX). INPP4B-CaaX dephosphorylates the D-4 position of phosphatidylinositol 3,4-bisphosphate and inhibits phagocytosis. One benefit is that generating stable cell lines is substantially faster than selecting for random integrations. Without FACS, the method typically gives ~50% of the cells that are transfected; with sorting, this approaches 100%. This makes phagocytosis experiments easier since more cells can be analyzed per experiment, allowing for population-based measurements where a ~10% transient transfection rate is insufficient. Finally, using the doxycycline-promoter allows for low near endogenous expression of proteins or robust overexpression.
Key features
• This protocol builds on the protocols and reagents developed by Kowarz et al. [1] and extends it to using RAW macrophages.
• Allows for the rapid generation of stably induced cell lines.
• This protocol also determines the phagocytic index and efficiency.
Keywords: Phagocytosis Phosphoinositides Stable inducible cells Sleeping beauty transposon system RAW264.7 Electroporation
Graphical overview
Protocol overview. Through electroporation or lipofection, RAW 264.7 cells are transfected with two plasmids: one carrying the Sleeping Beauty transposase and the other with an integration cassette carrying the gene of interest, reverse doxycycline-controlled repressor gene, and a puromycin-resistance gene. Transfected cells are then selected with 2.5 μg/μL of puromycin for at least 5 days to select for a polyclonal mix of stably transfected cells. For experiments, cells are treated with doxycycline to induce the TagBFP-INPP4B-CAAX or the inactive control TagBFP-INPP4BC842A-CAAX. Cells are exposed to opsonized particles to induce phagocytosis.
Background
The introduction of plasmid DNA into mammalian cells has been an essential method to support cellular and molecular biology investigations. Typically, the introduction of plasmid DNA can be mediated by a few approaches, including electroporation, chemical transformation, polyfection using polymer-based transfection reagents, microinjection, and viral transduction. These methods vary in efficiency depending on cell type and scale. Another factor to consider is whether the researcher wants transient over-expression of a gene of interest or generation of stably transfected cell lines. The generation of a stable cell line is typically more time-consuming. Still, it can be advantageous if cells are refractory to lipofection or if the same cell lines will be used for several experiments. In our lab, we use the Sleeping Beauty transposon system to help facilitate the integration of DNA into the genome to support the generation of stable cell lines [2–4]. This transposon system has been widely used as a genetic engineering tool for the last decade. Indeed, we have used this system to generate RAW264.7 cells expressing the reverse tetracycline repressor (rTetR) for controllable induction of our protein of interest, TagBFP-INPP4B-CaaX [5]. As these cells are generally challenging to transfect transiently, this approach allows for the rapid creation of cell lines.
The protocol described here is an adaptation of Kowarz et al. [1]. Briefly, this system utilizes a plasmid (SB100x) encoding the Sleeping Beauty transposase and a second plasmid containing an integration cassette with a gene of interest, a selectable marker, flanked by inverted terminal repeats. Kowarz et al. [1] generated a variety of plasmids, available from Addgene, to allow researchers to create cells with constitutive or doxycycline-inducible genes, an array of drug-resistant markers (e.g., puromycin or hygromycin) and the option also to deliver the reverse tetracycline repressor.
Phagocytosis is an actin-driven process used when cells ingest particles greater than 0.5 mm, such as pathogens and apoptotic bodies [6–8]. Phagocytosis entails the extension of actin-rich pseudopods to surround the target and pull the particle into the cell, forming a nascent phagosome [9]. The extensive actin remodeling, polymerization, bundling, and depolymerization must be exquisitely controlled and temporally coordinated. Major regulators in this process are the metabolism and interconversion of phosphoinositides lipids [10]. For instance, the generation of phosphatidylinositol (3,4)-bisphosphate (PtdIns3,4P2) at the engagement site is crucial for extending pseudopods to envelop prey [5]. Mammalian cells contain at least two pathways and multiple enzymes capable of synthesizing PtdIns3,4P2; thus, the knockdown of individual enzymes or specific inhibitors often generates unsatisfying results [11]. Instead, we used a synthetic plasmid-borne enzyme inositol polyphosphate 4-phosphatase type IIB (INPP4B) produced as a chimera with TagBFP for visualization, as well as a C-terminal CaaX motif for prenylation and membrane targeting [12]. This chimeric protein catalyzes the dephosphorylation of the D-4 position of PtdIns3,4P2 in the plasma membrane regardless of the biosynthetic route [12].
We generate doxycycline-inducible RAW264.7 cells expressing BFP-INPP4B-CaaX and a second cell line expressing the catalytically inactive BFP-INPP4BC842A-CaaX that serves as a control. Following overnight treatment with doxycycline to induce the INPP4B-containing constructs, we conduct phagocytosis assays to determine the role of PtdIns3,4P2 [5]. Typically, our phagocytosis assays use IgG-opsonized sheep red blood cells or IgG-coated polystyrene beads as prey to assess Fcg receptor-mediated phagocytosis [13].
Many approaches are available to assess phagocytosis; most rely on fluorescent labels on the prey to quantify internalization and phagosome maturation. Flow cytometry analysis determines the overall ingestion of particles by measuring total fluorescence intensity/cell as a parameter for particle uptake [14]. Other methods use pH-sensitive fluorescent particles that become brighter in acidic conditions, such as in the phagolysosome, that measure particle uptake and phagosome maturation [15]. However, both these approaches are limited in defining particle engagement vs. complete engulfment. Instead, we prefer a high-resolution confocal microscope and an antibody staining approach to distinguish bound vs. fully internalized particles (Figure 1).
Figure 1. INPP4B-CaaX inhibits the internalization of IgG opsonized particles. A. Representation of the assay. All sheep red blood cells (SRBCs) are labeled with the AF488-conjugated antibody, whereas only extracellular SRBCs are stained with the AF647 antibody at the end of the incubation period. B. Representative image. Fluorescence markers are pseudocolored: AF488 (green), AF647 (magenta), and BFP (cyan). Scale bar = 5 μm. C. A double y-axis graph depicting the results of panel B. The phagocytic index represents the number of beads internalized by the cells over a fixed period. Phagocytic efficiency represents the percentage of engaged particles that are internalized.
Materials and reagents
Biological materials
1. RAW264.7 cell line (American Type Culture Collection, catalog number: TIB-71)
2. Sheep red blood cells, 10% suspension (MP Biomedicals, catalog number: 55876), stored at 2–8 °C
3. pCMV(CAT)-T7-SB100X plasmid (Addgene, plasmid #34879)
4. pSBtet-Pur plasmid (Addgene, plasmid #60507)
5. Primary antibodies (store at -20 °C):
a. Anti-sheep red blood cell rabbit IgG antibody (85 mg/mL) (Rockland, catalog number: 113-4139)
b. Total human serum IgG antibody (50 mg/mL) (Millipore Sigma, catalog number: I4381)
6. Secondary antibodies (store at -20 °C):
a. Goat anti-rabbit IgG (H+L) antibody-AlexaFluor647 (Jackson ImmunoResearch, catalog number: 111-605-144)
b. Goat anti-rabbit IgG (H+L) antibody-AlexaFluor488 (Jackson ImmunoResearch, catalog number: 111-545-144)
c. Donkey anti-human IgG (H+L) antibody-AlexaFluor647 (Jackson ImmunoResearch, catalog number: 709-605-149)
d. Donkey anti-human IgG (H+L) antibody-AlexaFluor488 (Jackson ImmunoResearch, catalog number: 709-545-149)
Reagents
1. Paraformaldehyde 16% wt/vol (PFA) (Electron Microscopy Sciences, catalog number: 15700)
2. Doxycycline hyclate (Sigma-Aldrich, catalog number: D9891)
3. Puromycin (Sigma-Aldrich, catalog number: P8833
4. Roswell Park Memorial Institute Medium (RPMI) 1640 with L-glutamine and sodium pyruvate (Wisent Bioproducts, catalog number: 350-015-CL)
5. Phosphate buffered saline (PBS) without calcium and magnesium (Wisent bioproducts, catalog number: 311-013-CL)
6. Phosphate buffered saline with calcium and magnesium (PBS+/+) (Wisent bioproducts, catalog number: 311-011-CL)
7. Fetal bovine serum (FBS) (Wisent Bioproducts, catalog number: 080-150)
8. 4.2 mm Polystyrene beads with 2% divinylbenzene (Bangs Laboratories, Inc., catalog number: PS06005), stored at 2–8 °C
9. Fugene HD (Promega, catalog number: E2311)
10. Electrolytic buffer E2 (Invitrogen, catalog number: MPK10096E)
11. Resuspension buffer R (Invitrogen, catalog number: MPK10096R)
Solutions
1. 4% PFA (see Recipes)
2. Selection medium (see Recipes)
3. Induction medium (see Recipes)
Recipes
1. 4% PFA
Reagent Final concentration Amount
16% PFA 4% 1 mL
PBS+/+ n/a 3 mL
Total n/a 4 mL
2. Selection medium (2.5 μg/mL of puromycin in RPMI)
Reagent Final concentration Amount
RPMI 1640 n/a ~10 mL
Puromycin (10 mg/mL) 2.5 μg/mL 2.5 μL
Total n/a 10 mL
3. Induction medium (1 μg/mL of doxycycline in RPMI)
Reagent Final concentration Amount
RPMI 1640 n/a ~10 mL
Doxycycline hyclate (10 mg/mL) 1 μg/mL 2.5 μL
Total n/a 10 mL
Laboratory supplies
1. T-25 (Sarstedt, catalog number: 83.3910)
2. T-75 (Sarstedt, catalog number: 83.3911)
3. Cell scraper (Wuxi NEST Biotechnology, catalog number: 710001)
4. 6-well plates (Sarstedt, catalog number: 83.3920)
5. 12-well plates (Sarstedt, catalog number: 83.3921)
6. 18 mm circular cover glass, #1½ (Electron Microscopy Sciences, catalog number: 72222-01
7. Kimwipes (Kimberly-Clark ProfessionalTM 34155, catalog number: 06-666A)
8. 1.5 mL Eppendorf tubes (Froggabio, catalog number: LMCT1.7B)
9. 15 mL tubes (Fisher Scientific, catalog number: 14-959-53A)
Equipment
1. Incubator (Eppendorf, model: CellXpert C170, catalog number: 6734)
2. Tabletop centrifuge (Thermo Scientific, model: Pico 21, catalog number: 75002553)
3. Plate centrifuge (Thermo Scientific, model: Sorvall Legend RT, catalog number: 75004377)
4. Tube revolver (Thermo Scientific, catalog number: 88881001)
5. Vortex (Scientific Industries, catalog number: SI-0236)
6. Neon Invitrogen transfection system (Thermo Fisher Scientific, Invitrogen, catalog number: MPK5000)
7. Hemocytometer (NanoEntek, catalog number: EVE-MC)
8. 3i MarianasTM spinning-disc confocal microscopy, based on the Zeiss Axio Observer 7 Advanced Microscope with Definite Focus 3 [Intelligent Imaging Innovation (3i), custom-built]
Software and datasets
The microscope and image acquisition were controlled with SlideBook 2024 (3i). TIFF images were exported and analyzed in ImageJ2 Version 2.14.0/1.54f [16]. Representative images were chosen based on a good signal-to-noise ratio. Merging and cropping fluorescent channels were performed in ImageJ2. To aid visualization, linear adjustments were made to brightness and contrast across the entire image. To help with data accessibility and enhance the presentation of micrographs, we switched the default red lookup table to magenta. Graphical overview was generated using BioRender.com. Figure 1 was assembled using Adobe Illustrator 2024 and GraphPad Prism V9.
Procedure
A. Cell culture
1. Grow RAW 264.7 cells in tissue culture flasks (T-25 or T-75) in 10 mL or 20 mL of RPMI 1640 supplemented with 5% heat-inactivated FBS in an incubator at 37 °C under 5% CO2.
2. Wash cells twice with 3–5 mL of prewarmed PBS.
3. Aspirate PBS and gently scrape cells with a sterile cell scraper.
4. Add 5 mL of RPMI 1640 + 10% FBS and gently pipette cells into a homogenous suspension.
5. Transfer cells into a 15 mL conical tube and centrifuge at 500× g for 5 min.
6. Aspirate the medium without disturbing the pellet.
7. Resuspend in 10 mL of RPMI 1640 + 5% FBS and pipette gently to break cell clumps.
8. Every three days, split in 1:10 dilution.
B. Electroporation (Neon transfection system)
1. Prepare a 6-well plate with 2 mL of RPMI without antibiotics. Use one well per electroporation (e.g., 1 for INPP4B-CaaX and 1 for the catalytically inactive INPP4BC842A-CaaX).
2. Place in an incubator at 37 °C under 5% CO2 for at least 30 min.
3. Transfer grown cells (5–10 mL) into a 15 mL conical tube and count them using a hemocytometer. Ideally, there should be 1–2 × 106 cells per milliliter.
4. Centrifuge the cell suspension in a 15 mL conical tube at 500× g for 5 min.
5. Aspirate media and wash with 1 mL of PBS.
6. Centrifuge at 500× g for 5 min.
7. Set up a Neon tube with 3 mL of electrolytic buffer E2.
8. Set the pulse condition at 1,700 V for 20 ms on the Neon system.
9. Aspirate the PBS and resuspend the cell pellet in resuspension buffer R to reach a final density of ~1.0 × 107 cells per milliliter.
10. Transfer 100 μL of cells in resuspension buffer R to a new Eppendorf tube.
11. Add 1 μg of pCMV(CAT)-T7-SB100 and 10 μg of pSBtet-Pur-TagBFP2-INPP4B-CAAX plasmids into the Eppendorf tube containing cells resuspended in resuspension buffer R. Gently mix by pipetting. Ideally, the volume of DNA should be no more than 10 μL.
12. Remove the pre-incubated 6-well plate and place it in the biosafety cabinet.
13. Using a 100 μL Neon tip, draw from the DNA–cell solution and dock it appropriately in the Neon transfection system.
14. Run the program. After receiving the “Complete” message, immediately transfer the contents of the 100 μL tip to one well of the 6-well plate.
15. Place the plate overnight in the incubator at 37 °C under 5% CO2.
C. Transfection (Fugene HD)
1. Seed approximately 5 × 104 cells from Section A in each well of a 12-well plate overnight.
2. Add 1 μg of DNA in 100 μL of serum-free RPMI.
3. Add 3 μL of Fugene HD into the DNA–DMEM suspension and incubate for 15 min.
4. Distribute the mixture evenly into two wells (50 μL in each) and place it in an incubator overnight.
D. Stable cell line generation with the Sleeping Beauty transposon system
1. Electroporate or transfect cells with pCMV(CAT)-T7-SB100 and pSBtet-Pur-TagBFP2-INPP4B-CAAX or pSBtet-Pur-TagBFP2-INPP4B(C842A)-CAAX plasmids in a 1:10 ratio (at least 1 μg:10 μg for electroporation)
2. Replace media with 2 mL of selection media per sample for at least 5 days.
a. To enhance selection, replace old media with fresh puromycin-RPMI media after 2 days.
3. Transfer to T-25 in fresh media without puromycin to grow.
E. Sheep erythrocyte opsonization
1. Gently vortex the 10% red blood cell suspension.
2. Draw 100–200 μL of the suspension into an Eppendorf tube.
3. Centrifuge cells at 3,000× g for 30 s and aspirate the supernatant.
4. Wash with 1 mL of PBS.
5. Centrifuge cells at 3,000× g for 30 s and aspirate the supernatant.
6. Resuspend in 100 μL of PBS with 3 μL of anti-sheep red blood cell rabbit antibody (see Troubleshooting).
7. Incubate at 37 °C for 1 h in a shaking block heater at 300–400 rpm.
F. Polystyrene bead opsonization
1. Add 100 μL of PBS in an Eppendorf tube.
2. Gently vortex the polystyrene bead solution.
3. Transfer 20 μL of beads into the Eppendorf tube with PBS.
4. Add 20 μL of human IgG (50 mg/mL) to the bead–PBS solution.
5. Incubate at 37 °C for 1 h in an end-over-end rotator.
G. Phagocytosis assay
1. Seed ~5 × 104 of stable cells (expressing INPP4B-CAAX or INPP4B(C842A)-CAAX) on 18 mm coverslips in a 12-well plate.
2. Once adhered, add 1 mL of induction media to stable cells and incubate at 37 °C under 5% CO2 overnight.
3. Label IgG opsonized particles with 2 μL of the AF488-fluorescent antibody (goat anti-rabbit for red blood cells and donkey anti-human for polystyrene beads).
4. Incubate for 5 min at room temperature in an end-over-end rotator.
5. Centrifuge at 3,000× g for 2 min and aspirate the supernatant.
6. Wash with 1 mL of PBS and centrifuge at 3,000× g for 2 min.
7. Aspirate supernatant and resuspend with 100 μL of PBS.
8. Add 10, 20, or 40 μL (1×, 2×, 4×) of the opsonized particles to cells on a 12-well plate.
9. Centrifuge the 12-well plate at 1,000× g for 1 min.
10. Incubate at 37 °C under 5% CO2 for 10–15 min.
11. Aspirate media and wash with 500 μL of ice-cold PBS+/+.
12. Aspirate and add 500 μL of PBS with 1 μL of AF647-fluorescent antibody to label non-internalized particles.
13. Mix gently and incubate for 5–10 min.
14. Aspirate and wash with ice-cold PBS+/+.
15. Add 800 μL of 4% PFA and cover the plate with foil for 30 min to fix cells on the coverslip.
16. Wash fixed cells twice with 500 μL of PBS.
17. The plate can be stored at 4 °C for five days before microscopy.
H. Microscopy
The Fairn lab spinning-disc confocal microscope is provided by Intelligent Imaging Innovation (3i), Denver, Colorado. The 3i MarianasTM spinning-disc confocal microscopy is based on the Zeiss Axio Observer 7 Advanced Microscope with Definite Focus 3. Epifluorescence viewing and imaging of samples used pE-340Fura Illumination System (CoolLED) with GFP or mCherry filter cubes (Chroma). The system incorporates a Yokogawa CSU-W1 T2 super-resolution spinning disk confocal, 50 μm and SoRa disks with a magnification changer (1×, 2.8×, and 4×) for confocal and super-resolution imagining. The system has a Plan-Apochromat 20×/0.8 NA and a C Plan-Apochromat 63×/1.4 NA oil objective (Zeiss). The system uses four lasers controlled by a LaserStack v4 with single-mode optical fibers for 405 nm (150 mW), 488 nm (200 mW), 561 nm (140 mW), and 638 nm (200 mW). Samples were collected using appropriate single bandpass filters or a quad-band filter (440/521/607/700 nm). Images were acquired with a Hamamatsu ORCA Fusion BT sCMOS camera with 2,304 × 2,304 pixels and a pixel size of 6.5 μm controlled with SlideBook software (3i). Acquisition settings and capture were controlled by SlideBook Software v2024 (Intelligent Imaging Innovations).
Data analysis
Post-acquisition images were analyzed, and the number of internal (single color) and external/bound particles were counted. Typical results are illustrated in Figure 1. For experiments, we usually compare uninduced to induced samples or, where possible, active enzyme vs. inactive enzyme. The results are presented as the mean ± standard error of n = 3–5 experiments, with 100 technical experiments (cells) per individual condition. Student’s t-test (2 groups) or Analysis of Variance (ANOVA) (> 2 groups) with an appropriate post-hoc test such as Tukey’s honest significance difference are used to test significance.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article:
• Montaño-Rendón et al. [5]. PtdIns(3,4)P2, Lamellipodin, and VASP coordinate actin dynamics during phagocytosis in macrophages. Journal of Cell Biology (Figure 3A and B).]
General notes and troubleshooting
General notes
1. Use early passage cells when making stable cell lines.
2. Determine the optimal doxycycline concentration for the desired level of expression. If robust over-expression is required, 1 mg/mL of doxycycline for 18 h is typically a good starting point. However, if the goal is to express genes of interest at near endogenous levels, we suggest using 100 ng/mL.
3. The protocol described is for RAW264.7 cells; however, we have also used this approach with other cell types, such as HeLa [17], HCT116 [18], and ARPE19 [19].
4. The protocol generates a polyclonal cell population. We prefer this approach in most cases since it prevents potential issues with expanding individual clones. However, following the introduction of the plasmids into the RAW264.7 cells, they can be diluted, and single cells can be aliquoted in each well of a 96-well plate for selection and expansion. RAW cells grow better in these situations when fresh media is supplemented with 2-day-old conditioned media (50:50).
5. In this example, we used spinning disc confocal microscopy. However, this assay can be done with less sophisticated systems. In our lab, we also use an EVOS M5000 widefield imaging system, which illuminates samples with three specific LEDs.
Troubleshooting
Opsonization of sheep red blood cells may result in agglutination, which can be observed as clumps after the incubation. To avoid agglutination, each vial of rabbit anti-sheep red blood cell antibody should be titrated (2–5 mL per milliliter).
Bangs Laboratories also sells latex beads without 2% DVB. In our experience, the human IgG does not adhere to these particles.
Phagocytosis is temperature-dependent. Ensure the media is prewarmed and cells are maintained at 37 °C during the assay.
Acknowledgments
This work was supported by a Project Grant from the Canadian Institutes of Health Research (PJT165968) to G.D.F. This bio-protocol is based on a previous publication from the laboratory, Montaño-Rendón et al. [5] Journal of Cell Biology (2022) 221(11): e202207042 and Cabral-Dias et al. [19] Journal of Cell Biology (2022) 221(4): e201808181.
Competing interests
The authors declare no competing interests.
References
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Montaño-Rendón, F., Walpole, G. F., Krause, M., Hammond, G. R., Grinstein, S. and Fairn, G. D. (2022). PtdIns(3,4)P2, Lamellipodin, and VASP coordinate actin dynamics during phagocytosis in macrophages. J Cell Biol. 221(11): e202207042.
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Article Information
Publication history
Received: Sep 30, 2024
Accepted: Dec 2, 2024
Available online: Dec 26, 2024
Published: Feb 5, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Immunology > Immune cell function > Macrophage
Cell Biology > Cell-based analysis > Gene expression
Immunology > Immune mechanisms
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Original Research Article:
The authors used this protocol in The Journal of Biological Chemistry May 2023
Abstract
Amylin is an amyloidogenic neuroendocrine hormone co-synthesized and co-secreted with insulin from the pancreas. It readily crosses the blood–brain barrier and synergistically forms mixed amyloid plaques with β-amyloid (Aβ) in brain parenchyma. Parenchymal amylin-Aβ plaques are found in both sporadic and early-onset familial Alzheimer’s disease (AD), yet their (patho)physiological role remains elusive, particularly due to a lack of detection modalities for these mixed plaques. Previously, we developed an enzyme-linked immunosorbent assay (ELISA) capable of detecting amylin-Aβ hetero-oligomers in brain lysate and blood using a polyclonal anti-amylin antibody to capture hetero-oligomers and a monoclonal anti-Aβ mid-domain detection antibody combination. This combination allows for the recognition of distinct amylin epitopes, which remain accessible after amylin-Aβ oligomerization has begun, and precise detection of Aβ epitopes available after oligomer formation. The utility of this assay is evidenced in our previous report, wherein differences in hetero-oligomer content in brain tissue from patients with and without AD and patients with and without diabetes were distinguished. Additionally, using AD model rats, we provided evidence that our assay can be employed for the detection of amylin-Aβ in blood. This assay and protocol are important innovations in the field of AD research because they meet an unmet need to detect mixed amyloid plaques that, if targeted therapeutically, could reduce AD progression and severity.
Key features
• Detects amylin-Aβ hetero-oligomers in blood from patients with Alzheimer’s disease.
• Enables simultaneous, high-throughput analysis of hetero-oligomer content of brain and blood tissue.
• Allows exploration into the amylin-Aβ interaction during AD pathogenesis, potentially leading to novel treatment mechanisms by controlling the amylin-Aβ interaction.
Keywords: Amylin Amyloid Alzheimer’s disease Beta-amyloid Enzyme-linked immunosorbent assay Hetero-oligomers
Graphical overview
Detection of amylin-Aβ hetero-oligomers by ELISA. (A) Amyloidogenic amylin and Aβ form mixed hetero-oligomers in brain tissue. (B) The ELISA described in this protocol is capable of detecting these mixed hetero-oligomers. (C) In the future, perhaps the detection of these mixed hetero-oligomers can assist in refining diagnostic measures and treatments for Alzheimer’s disease.
Background
Alzheimer's disease (AD) has previously been linked to dysregulated pancreatic hormones, including insulin, which can exacerbate AD pathology by causing brain insulin resistance [1–2]. Amylin, a centrally acting neuroendocrine hormone co-synthesized and co-secreted with insulin from pancreatic β-cells [3], readily crosses from the blood to the brain parenchyma [4] and is reportedly involved in the central regulation of satiation [5]. In nearly all (>95% prevalence) patients with type-2 diabetes mellitus (T2DM), amylin forms cytotoxic amyloid plaques in the pancreas, which contributes to the pathogenesis of T2DM by inducing β-cell apoptosis and subsequent β-cell mass depletion [6–8]. In both sporadic and early-onset familial AD, amylin synergistically forms co-aggregates with vascular and brain parenchymal Aβ in the brain [9–14]. Consistent with findings in human AD brains, APPswe/PS1dE9 (APP/PS1) rats expressing human amylin in the pancreas (since murine amylin is non-amyloidogenic) revealed that chronic exposure to circulating amylin promoted cerebrovascular and parenchymal amylin-Aβ deposition [9–13]. These results suggest an association between Aβ pathology and amyloidogenic human amylin but the exact mechanism responsible for amylin-Aβ co-aggregation remains to be determined.
Previous research demonstrated that heterologous seeding between amylin and common Aβ fragments (e.g., Aβ42 and Aβ40) promotes amyloid formation in a manner that is comparable to that of homogenous amylin amyloid [15–17]. In vivo amylin-Aβ hetero-amyloid formation and cytotoxicity are facilitated by the co-expression of the two peptides in cells [21–22]. The hypothesis that amylin and Aβ peptides aggregate in the human AD brain to form amylin-Aβ hetero-oligomers has previously been verified by co-immunoprecipitation experiments using human brain samples from AD patients [9]. These converging data [9,15–22] support the presence of amylin-Aβ hetero-oligomers in the human brain and establish a need for their quantification. Proper detection and quantification of amylin-Aβ hetero-oligomers may further establish the connection between amylin and Aβ and their collective role in AD pathology.
While the need for a means to reliably quantify the molecular amylin-Aβ interaction in AD has been acknowledged [9–13], currently employed molecular techniques (namely immunoprecipitation, western blot, circular dichroism, and electron microscopy) limit studies to reduced sample sizes due to their inherent complexity, non-scalability, and in some cases, cost. Additionally, the denaturing conditions commonly used in methods such as western blot prove to be additional obstacles, as the size and structural stability of hetero-oligomers can be altered. Therefore, there is still an unmet need for an assay method that can detect and quantify hetero-oligomers of amylin-Aβ without interrupting molecular interactions. Previously, we reported the creation of a novel enzyme-linked immunosorbent assay (ELISA) to detect amylin-Aβ hetero-oligomers in blood and brain tissue [23] in AD model animals and human donor tissue. Here, we provide a detailed, stepwise protocol for our assay. Relative to the currently employed techniques, this assay is scalable, high-throughput, cost-effective, and non-complex, all while critically maintaining hetero-oligomer stability. The assay detailed here may prove to be a useful tool to detect, analyze, and eventually disentangle the role of the amyloidogenic amylin and Aβ proteins in AD and other pathological conditions.
Materials and reagents
Biological materials
1. Mouse anti-human-Aβ(1-16) antibody (BioLegend, catalog number: 803002)
2. Rabbit anti-amylin P2 antibody (not commercially available, see [23])
3. Mouse anti-human-total-Aβ (BioLegend, catalog number: 800720)
4. Aβ40 synthetic peptide (Anaspec, catalog number: AS-24235)
Reagents
1. ELISA assay diluent (5×) (BioLegend, catalog number: 421203)
2. Dimethyl sulfoxide (DMSO) (Sigma, catalog number: D8418)
3. Na2CO3 (Sigma, catalog number: S7795)
4. NaHCO3 (Sigma, catalog number: S5761)
5. 10× PBS (Corning, catalog number: 46-013-CM)
6. 10% Tween-20 (Bio-Rad, catalog number: 1610781)
7. Aβ40 synthetic peptide (Anaspec, catalog number: AS-24235)
8. 3,3',5,5' tetramethylbenzidine (TMB) substrate (Thermo Scientific, catalog number: 34028)
9. Stop solution (Thermo Scientific, catalog number: N600)
Solutions
1. Bicarbonate buffer (see Recipes)
2. PBST washing buffer (see Recipes)
Recipes
1. Bicarbonate buffer (250 mL)
Note: Adjust the solution’s pH to 9.6. Use distilled water as diluent.
Reagent Final concentration Mass
Na2CO3 0.028 M 0.7575 g
NaHCO3 0.071 M 1.50 g
2. PBST washing buffer (500 mL)
Note: Dilute 50 mL of 10× PBS to 1× using 450 mL of distilled water as diluent before preparing PBST washing buffer.
Reagent Final concentration Volume
1× PBS - 500 mL
10% Tween-20 0.05% 2.5 mL
Laboratory supplies
1. 96-well clear flat-bottom polystyrene high bind microplate (Corning, catalog number: 9018)
2. Sealing film, polyester (VWR, catalog number: 89134-430)
3. Absorbent paper towels (Pacific Blue Basic, catalog number: 23504)
4. epT.I.P.S. standard 20–300 μL pipette tips (Eppendorf, catalog number: 022492047)
5. 1.5 mL standard line microcentrifuge tubes (VWR, catalog number:10025-726)
6. 50 mL reagent reservoirs (Corning, catalog number: 4870)
7. 8-channel 30–300 μL multichannel pipette (USA Scientific, catalog number: 7108-3300)
Equipment
1. xMark microplate absorbance spectrophotometer (Bio-Rad, catalog number: 1681150)
Software and datasets
1. Microsoft Excel (2019)
2. Microplate Manager Software 6 (MPM) (Bio-Rad, v6.3)
Procedure
This protocol has been validated using frozen and freshly prepared samples. The homogenization buffers and protocol used previously are described in [23]. Preparation of samples in other buffers or using other homogenization methods may be possible so long as the conditions do not disrupt amylin-Aβ binding interactions.
Unless otherwise noted, bring all reagents to room temperature prior to use. Determine ELISA plate layout/template prior to use; we recommend using the first two columns (16 wells) for the standard curve (8 standards in duplicate). Dilute 5× assay diluent to 1× final concentration using 1× PBS prior to use. A multichannel pipette is used for the addition of all solutions, except the standards/controls, which are added individually. A graphical summary of the protocol is shown in Figure 1.
Figure 1. Graphical summary of ELISA protocol. General overview of the present ELISA protocol, highlighting key steps. Note that after the addition of the stop solution, the plate must be read in a spectrophotometer, which is not depicted in this summary figure. Data analysis steps are not shown.
A. Coating with capture antibodies
1. Dilute the mouse anti-human-Aβ(1-16) antibody to a final dilution of 1:400 using bicarbonate buffer and coat the wells of the 96-well plate to be used for the standards with 100 μL.
2. Dilute the rabbit anti-amylin P2 antibody to a final dilution of 1:400 using bicarbonate buffer and coat all sample wells of the plate with 100 μL.
3. Seal the plate with sealing film and incubate overnight at 4 °C without shaking.
B. Loading standards and samples
1. Prepare the standards.
a. Resuspend the Aβ40 peptide in a sufficient amount of DMSO to prepare the 1 ng/mL stock solution.
b. Serial dilute the stock in 1× assay diluent to create the following concentrations (in ng/mL): 10, 5, 2.5, 1.25, 0.625, 0.3125, 0.1562. Use assay diluent buffer as a blank (“0”) standard.
2. Remove the plate sealer and decant antibody solutions by gently shaking the plate (upside down) over a sink. Remove any residual solution by gently tapping the plate on an absorbent paper towel.
a. When tapping the plate, tap hard enough that any residual solution is removed from the wells and the wells are dry.
b. Repeat this method of washing/tapping for all following solution decanting steps.
3. Wash the plate with 300 μL of PBST washing buffer per well. Remove any remaining wash buffer by tapping the plate on an absorbent paper towel, as above. An automatic plate washer may also be used if available.
4. Add 300 μL of 1× assay diluent to each well and incubate for 1 h at room temperature.
5. Decant blocking solution and remove any residual solution as described above.
6. Wash the plate with 300 μL of PBST washing buffer per well 1–2 times. Decant and tap after each wash, as described above.
7. Add 100 μL of the Aβ40 peptide standard to the appropriate wells (coated with the anti-human-Aβ(1-16) antibody only). Include 100 μL of 1× assay diluent as the “0” standard/blank, as described above.
8. Dilute samples to 1:2 final dilution using 1× PBS for a final volume of 200 μL (100 μL per well, in duplicate). Plate 100 μL of diluted sample to the appropriate wells (coated with the anti-amylin P2 antibody).
9. Add 100 μL of 1× assay diluent as a blank for the samples (for wells originally coated with the P2 antibody).
10. Seal the plate and incubate overnight at 4 °C.
C. Developing the assay and data collection
1. Prepare the HRP-conjugated anti-human-total-Aβ detection antibody by diluting to a final dilution of 1:400 in 1× assay diluent.
2. Remove the plate sealer, decant the standards and samples, and remove any remaining solution, as above.
3. Wash the plate with 300 μL of PBST washing buffer per well 3 times. Decant and tap after each wash, as above.
4. Add 100 μL of prepared detection antibody per well and incubate at room temperature for 1 h.
a. It is recommended to remove the TMB from 4 °C at this time, to allow it time to acclimate to room temperature before use.
5. Decant the antibody and remove the residual solution as above.
6. Wash the plate with 300 μL of PBST washing buffer 3–4 times with decanting and tapping.
7. Add 100 μL of TMB substrate (at room temperature) per well. Incubate for approximately 30 min until standard wells show gradation and sample wells show a signal. Check the plate after 20 min of incubation and then every few minutes to prevent oversaturation of signal intensity.
8. Add 50 μL of stop solution per well in the exact same order in which the substrate was added.
9. Read the plate at 450 nm in a spectrophotometer.
Data analysis
Following data acquisition using MPM6 software, data are exported to Excel for further analysis. In each experiment, samples should be run in duplicate or triplicate for technical replicates. Average the values for all duplicate/triplicate wells. Subtract the average absorbance value of the standard blank wells (the wells with only 1× assay diluent coated with anti-human-Aβ(1-16) antibody in step A1) from all standard wells. Subtract the average absorbance value of the sample blank (the wells with only 1× assay diluent coated with anti-amylin P2 antibody in step A1) from all sample wells to remove background and non-specific signal. Create a standard curve using the absorbance values vs. standard concentrations and generate the linear regression equation. The concentration of samples can then be interpolated from the standard curve regression line. A detailed description of the data analysis method appears in our previous report [23] in the “Experimental procedures” section. An overview of the data analysis method is shown in Figure 2.
Figure 2. Data analysis overview. After exporting the values to Excel, values for duplicate wells are averaged together. For all standard values, the average of the “0” standard is subtracted. For all samples, the “1× assay diluent” average is subtracted. Then, the standard values are used to create a standard curve and generate a linear regression equation, which can be used to calculate sample values.
Validation of protocol
This protocol has been used and validated in the following research article:
• Kotiya et al. [23]. Rapid, scalable assay of amylin-β amyloid co-aggregation in brain tissue and blood. Journal of Biological Chemistry (Figure 1, panels E–F; Figure 2 panels A–E; Figure 3, panels B–E; Figure 4 panels A–C; Figure 6 panels A–E).
General notes and troubleshooting
General notes
1. For convenience, we have prepared an example plate layout (Table S1) showing standards and samples plated in duplicate. We have also included a blank plate layout, which can be filled out with sample identifiers (Table S2).
2. The inherent instability and amyloidogenicity of both amylin and β-amyloid can cause variability between measurements. We strongly recommend that all samples that are to be compared against one another be run on the same assay plate. If necessary, samples can be run as singlets in two separate plates, and the values averaged together.
3. Following the acquisition of concentrations of samples (in ng/mL), we recommend normalizing each value to the respective sample’s total protein concentration to account for individual differences and variances that occur as a result of sample preparation. We recommend reporting final values as “ng amylin/mg total protein” (or similar).
4. Though not required, we recommend wrapping the plate with aluminum foil to prevent light interference and smudging/fingerprinting on the plate’s clear bottom, which would interfere with absorbance readings.
5. Placing the plate on a plate shaker during any incubation step is not recommended; conduct all incubation and washing steps without shaking.
6. When adding solutions, we recommend always adding solutions (wash buffer, TMB, stop solution, etc.) from top to bottom, ensuring duplicate/triplicates receive each solution simultaneously.
7. We recommend initially diluting samples two-fold with 1× PBS, as described in the protocol above. Samples may need to be run with higher (if the signal is too strong) or lower (if the signal is too weak) dilution factors according to amylin-Aβ concentration in the prepared sample.
8. Although Aβ40 does not form amyloid by itself rapidly [24], we recommend aliquoting Aβ40 stock to prevent frequent freeze/thaw cycles, which can impact aggregation formation.
Troubleshooting
Problem 1: Oversaturation of standards or samples.
Possible cause: Insufficient number of washing and/or decanting steps.
Solution: Modify the number of washes during sections B and C and ensure proper decanting to remove excess/residual solution from each well before adding the next solution.
Problem 2: No (or little) gradation between standards.
Possible cause: Multiple freeze/thaw cycles of Aβ standards, incorrect preparation, or dilution.
Solution: Minimize the number of freeze/thaw cycles of the Aβ standards by aliquoting stock peptide. Only thaw the peptide standard when prepared to run the assay. Pipette the standards carefully and ensure no bubbles are present when loading the standards in the wells. If necessary, run repeat experiments with the standard curve plated in triplicate.
Supplementary information
The following supporting information can be downloaded here:
1. Table S1. Example plate layout
2. Table S2. Blank plate layout
Acknowledgments
F.D. acknowledges the following funding sources: National Institutes of Health R01 NS116058, R01 AG057290, R01 AG053999.
This protocol was adapted and modified from Kotiya et al. [23].
Competing interests
The authors declare no competing interests.
References
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Article Information
Publication history
Received: Aug 8, 2024
Accepted: Dec 5, 2024
Available online: Dec 26, 2024
Published: Feb 5, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Neuroscience > Nervous system disorders > Neurodegeneration
Neuroscience > Basic technology > High-throughput screening
Biochemistry > Protein > Immunodetection
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Isolation of Intact Mitochondria From Drosophila melanogaster and Assessment of Mitochondrial Respiratory Capacity Using Seahorse Analyzer
CG Christopher M. Groen
AW Anthony J. Windebank
Published: Vol 15, Iss 3, Feb 5, 2025
DOI: 10.21769/BioProtoc.5180 Views: 34
Reviewed by: Darrell CockburnAnnmary Paul Erinjeri Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in The Journal of Neuroscience Feb 2022
Abstract
Analysis of mitochondrial function has broad applicability in many research specialties. Neurodegenerative disorders such as chemotherapy-induced peripheral neuropathy (CIPN) often exhibit damaged mitochondria or reduced mitochondrial respiratory capacity. Isolation of intact mitochondria for protein analysis or respiration measurements has been previously reported in numerous model organisms. Here, we describe an adaptation of previous protocols to isolate intact functional mitochondria from Drosophila melanogaster for use in a model of CIPN. Whole Drosophila are ground in isolation buffer, and mitochondria are purified using differential centrifugation through a sucrose and mannitol solution. The intact mitochondria are plated as a monolayer for measurements of mitochondrial oxygen consumption rates and response to inhibitor compounds on an Agilent Seahorse analyzer. This experimental protocol is quick and yields a purified population of intact mitochondria that may be used for functional assays for several hours after isolation. The isolated mitochondria may be used for respiration measurements, which reflect their health, and stored for protein or genetic analysis. Mitochondrial populations from multiple strains or treatment groups can be easily compared simultaneously. The rapid biochemical assessment of mitochondria, in combination with the utility of Drosophila as an in vivo genetic model system, offers great potential for researchers to probe the impact of genetics and pharmacologic interventions on mitochondrial respiratory capacity.
Key features
• This protocol describes rapid isolation of intact, functional mitochondria that may be used for respiration measurements or other biochemical analyses.
• Mitochondria isolated from Drosophila are assessed in an Agilent Seahorse analyzer utilizing multiple substrates and electron transport chain inhibitors to fully characterize mitochondrial respiratory capacity.
• This protocol is optimized to use Drosophila for easy in vivo genetic and pharmacologic manipulation, and assessment of the impact on mitochondrial function.
Keywords: Mitochondria Respiration Drosophila Seahorse Differential centrifugation Oxygen consumption
Graphical overview
Created in BioRender.com
Background
Mitochondria function is increasingly recognized as an essential factor in human health and disease. Altered mitochondria function is observed in cancer [1], aging [2–4], metabolic disorders [5], and neurodegenerative disorders such as Parkinson’s Disease [6] and chemotherapy-induced neuropathy [7,8]. Assessment of mitochondrial function and cellular respiration rates can reveal disease states in an organism or tissue. These measurements are invaluable tools to parse the reactions of cells to acute toxic stressors, genetic manipulations, or potential preventive and therapeutic interventions [9]. Analysis of mitochondrial health has progressed from earlier techniques using Clark electrodes to measure O2 consumption in individual samples to more advanced instrumentation such as Agilent Seahorse analyzers, which are capable of simultaneous analysis of 96-well plates. Modern respiration analyzers can also measure real-time response to the addition of substrates, metabolites, and mitochondria inhibitors. Mitochondria respiration analysis has been measured using intact cells in cell culture plates, dissociated tissues, and isolated mitochondria. Insights gained from these experiments advance our understanding of the role of mitochondria in disease and provide new potential biomarkers for the assessment of disease states [9].
Previous publications have described detailed protocols for the measurement of cellular respiration [9–11], isolation of mitochondria for metabolic analysis [12], and use of isolated mitochondria from mouse muscle tissue for oxygen consumption rate analysis in Seahorse analyzers [13]. Drosophila-specific protocols in this field include detailed experimental methods for isolation of larval mitochondria [14], metabolic analysis of flight muscle mitochondria [12], and isolation of mitochondria for enzymatic analysis of electron transport chain components [15]. The protocol described here is a modification of previous mitochondria isolation procedures from Drosophila, with optimization of Seahorse analysis of these isolated mitochondria to assess differences in respiratory capacity between different fly strains treated with chemotherapy agents known to damage mitochondria. This protocol was previously described in a manuscript reporting differences in cisplatin sensitivity between Drosophila strains and showed that the mitochondria have different basal respiration rates and different abilities to maintain activity following treatment with toxic stressors like cisplatin [16].
The protocol described here allows for rapid isolation of a pure mitochondria pellet. The mitochondria retain their activity for several hours after isolation if kept on ice. In addition, the intact mitochondria may be used for protein analysis or enzymatic activity assays. Isolation of mitochondria allows for direct assessment of electron transport chain function and the impact of multiple substrates and inhibitors. This allows researchers to focus on specific aspects of cell metabolism without the variables presented by whole-cell metabolomics. The availability of substrates can be tightly controlled by the user, whereas whole-cell or tissue respiration measurements must consider the availability of metabolites contributed by the whole-cell environment. ADP is often used as an injection compound to measure mitochondrial respiration before and after the substrate for ATP synthase is available. Respiration through Complex I of the electron transport chain may be measured through the use of pyruvate and malate as initial substrates. Succinate can be used as a substrate for Complex II, and ascorbate is a substrate for Complex IV. Likewise, inhibitors block components of the electron transport chain. Rotenone and antimycin A block Complex I and Complex III, respectively. Oligomycin blocks the activity of ATP synthase, while FCCP uncouples oxygen consumption from ATP synthesis. Careful selection of substrates and inhibitors can reveal a great deal about the function of isolated mitochondria. This method does not, however, provide a complete picture of cellular respiration, as the mitochondria are removed from the context of the cell. The lack of whole cells limits data collection from the Seahorse to oxygen consumption rate (OCR). The extracellular acidification rate (ECAR) measured by the Seahorse as a readout of glycolytic activity is not useful when only assessing isolated mitochondria. Researchers should carefully choose an experimental protocol based on specific goals in characterizing cellular metabolism.
Materials and reagents
Biological materials
1. Drosophila melanogaster stocks (Bloomington Drosophila Stock Center, stock numbers 36303 and 36304 were used in the sample data)
Reagents
1. Sucrose (Sigma-Aldrich, catalog number: S1888-1KG)
2. D-Mannitol (Sigma-Aldrich, catalog number: M4125-100G)
3. HEPES buffer, 1 M, pH 7.2 (Corning, catalog number: 25-060-Cl)
4. EDTA, 0.5 M (Sigma-Aldrich, catalog number: 03690-100ML)
5. Bovine serum albumin (BSA) (Sigma-Aldrich, catalog number: 05470-5G)
6. Potassium hydroxide (KOH) (Sigma-Aldrich, catalog number: 221473-25G)
7. Monobasic potassium phosphate (KH2PO4) (Sigma-Aldrich, catalog number: 1551139-5G)
8. Magnesium chloride (MgCl2) (Sigma-Aldrich, catalog number: M8266-100G)
9. EGTA, 0.5 M (RPI, catalog number: E14100-50.0)
10. Pyruvic acid (Sigma-Aldrich, catalog number: 107360-25G)
11. Malic acid (Sigma-Aldrich, catalog number: M6413-25G)
12. Adenosine 5’-diphosphate monopotassium salt dihydrate (ADP) (Sigma-Aldrich, catalog number: A5285)
13. Oligomycin A (Sigma-Aldrich, catalog number: 75351)
14. Carbonyl cyanide 4-(trifluoromethoxy) phenylhydrazone (FCCP) (Sigma-Aldrich, catalog number: C2920)
15. Rotenone (Sigma-Aldrich, catalog number: R8875)
16. Antimycin A (Sigma-Aldrich, catalog number: A8674)
17. Seahorse XF calibrant (Agilent, catalog number: 100840-000)
18. DC Protein Assay Kit I (Bio-Rad, catalog number: 5000111)
19. 95% Ethanol (Millipore-Sigma, catalog number: 65348-M)
Solutions
1. 10% w/v BSA (see Recipes)
2. Pyruvate stock solution (see Recipes)
3. Malate stock solution (see Recipes)
4. ADP stock solution (see Recipes)
5. Oligomycin A stock solution (see Recipes)
6. FCCP stock solution (see Recipes)
7. Rotenone stock solution (see Recipes)
8. Antimycin A stock solution (see Recipes)
9. Mitochondria isolation buffer (see Recipes)
10. Mitochondria assay buffer (2× stock) (see Recipes)
11. Working mitochondria assay buffer (1×) with substrates (see Recipes)
Recipes
1. 10% w/v BSA
Reagent Final concentration Quantity or Volume
BSA 10% w/v 1 g
ddH2O n/a *
Total (optional) n/a 10 mL
*Fully dissolve BSA in ~5–7 mL of ddH2O and then bring the solution to a final volume of 10 mL.
Aliquot and store at -20 °C.
2. Pyruvate stock solution (100 mM)
Reagent Final concentration Quantity or Volume
Pyruvic acid 100 mM 0.8806 g
ddH2O n/a 100 mL
Total (optional) n/a 100 mL
Aliquot and store at -20 °C.
3. Malate stock solution (100 mM)
Reagent Final concentration Quantity or Volume
Malic acid 100 mM 1.34 g
95% ethanol n/a 100 mL
Total (optional) n/a 100 mL
Aliquot and store at -20 °C.
4. ADP stock solution (100 mM)
Reagent Final concentration Quantity or Volume
ADP 100 mM 5.0132 g
ddH2O n/a 100 mL
Total (optional) n/a 100 mL
Aliquot and store at -20 °C.
5. Oligomycin A stock solution (10 mM)
Reagent Final concentration Quantity or Volume
Oligomycin A 10 mM 0.079 g
95% ethanol n/a 10 mL
Total (optional) n/a 10 mL
Aliquot and store at -20 °C.
6. FCCP stock solution (10 mM)
Reagent Final concentration Quantity or Volume
FCCP 10 mM 0.025 g
95% ethanol n/a 10 mL
Total (optional) n/a 10 mL
Aliquot and store at -20 °C.
7. Rotenone stock solution (5 mM)
Reagent Final concentration Quantity or Volume
Rotenone 5 mM 0.019 g
95% ethanol n/a 10 mL
Total (optional) n/a 10 mL
Aliquot and store at -20 °C.
8. Antimycin A stock solution (10 mM)
Reagent Final concentration Quantity or Volume
Antimycin A 10 mM 0.055 g
95% ethanol n/a 10 mL
Total (optional) n/a 10 mL
Aliquot and store at -20 °C.
9. Mitochondria isolation buffer
Reagent Final concentration Quantity or Volume
Sucrose 70 mM 4.79 g
D-Mannitol 210 mM 7.65 g
1 M HEPES, pH 7.2 5 mM 1 mL
0.5 M EDTA 1 mM 0.4 mL
10% BSA 0.5% 10 mL*
KOH n/a pH to 7.2
ddH2O n/a **
Total (optional) n/a 200 mL
*BSA is included in the isolation buffer and assay buffers to bind free fatty acids released during tissue homogenization and helps to maintain mitochondrial membrane potential and respiratory function.
**Fully dissolve solid reagents in ~100 mL of ddH2O. Use KOH to bring the pH to 7.2 and then bring the solution to a final volume of 200 mL.
Aliquot and store at -20 °C
10. Mitochondria assay buffer (2× stock)
Reagent Final concentration Quantity or Volume
Sucrose 140 mM 9.58 g
D-Mannitol 440 mM 15.3 g
KH2PO4 20 mM 0.544 g
MgCl2 10 mM 0.19 g
1 M HEPES, pH 7.2 4 mM 0.80 mL
0.5 M EGTA 2 mM 0.80 mL
10% BSA 0.4% 8 mL
KOH n/a pH to 7.2
ddH2O n/a *
Total (optional) n/a 200 mL
*Fully dissolve solid reagents in ~100 mL of ddH2O. Use KOH to bring the pH to 7.2 and then bring the solution to a final volume of 200 mL.
Aliquot and store at -20 °C
11. Working mitochondria assay buffer (1×) with pyruvate and malate
Reagent Final concentration Quantity or Volume
Mitochondria assay buffer (2× stock) 1× 15 mL
100 mM pyruvate stock solution 10 mM 3 mL
100 mM malate stock solution 5 mM 1.5 mL
ddH2O n/a 10.5 mL
Total (optional) n/a 30 mL
Stock solutions and buffers should be aliquoted and stored at -20 °. Make this solution fresh on the day of the assay.
Laboratory supplies
1. 1.5 mL Eppendorf tubes (Sigma-Aldrich, catalog number: EP022363212)
2. 15 mL centrifuge tubes (Corning, catalog number: CLS430052)
3. 50 mL centrifuge tubes (Corning, catalog number: CLS430828)
4. Pestles (Sigma-Aldrich, catalog number: BAF199230000)
5. Seahorse FluxPak (Agilent)
Note: This protocol describes experiments performed with Seahorse XF24 FluxPaks. This model has been discontinued and replaced by the Seahorse XFe24 analyzer and FluxPaks (Agilent, catalog number: 102340-100)
6. 96-well clear flat bottom microplates (Corning, catalog number: 353072)
Equipment
1. Sorvall Legend X1R centrifuge (Thermo Scientific, catalog number: 75004220)
2. M-20 microplate swinging bucket rotor (Thermo Scientific, catalog number: 75003624)
3. Eppendorf Centrifuge 5425 (Sigma-Aldrich, catalog number: 5405000646)
4. SpectraMax M3 microplate reader (Molecular Devices, catalog number: M3)
5. CO2 tank (for Drosophila anesthesia)
6. CO2 regulator (for Drosophila anesthesia) (Flystuff, catalog number: 59-143)
7. CO2 fly pad (Flystuff, catalog number: 59-114)
8. CO2 blowgun (Flystuff, catalog number: 54-104)
9. Fly brushes (Flystuff, catalog number: 59-204)
10. Fisher Scientific Isotemp incubator, non-CO2 (Fisher Scientific, catalog number: 15-103-0514)
11. Seahorse XF24 analyzer (Agilent)
Note: The current equivalent model Agilent Seahorse is the XFe24. Any Agilent Seahorse will be able to measure oxygen consumption from isolated mitochondria. Volumes and concentrations of reagents should be optimized for the analyzer.
Software and datasets
1. WAVE Desktop v2.6 (Agilent, 2018)
2. Prism v10.3.1 (GraphPad, 2024)
3. Microsoft Excel (Office 365, 2024)
Procedure
A. Drosophila melanogaster preparation
Note: Preparation of Drosophila may require 1–2 weeks or more to reach the desired age or complete treatment protocols. Plan experiments accordingly.
1. Collect adult Drosophila of the desired age(s) and genotype(s) in fresh food vials.
a. Ensure uniform age of Drosophila by removing all adult flies from adults-producing vials the day before collection of experimental flies.
b. Collect experimental flies in new food vials.
c. Maintain consistent numbers in each vial so that fly density does not influence experimental outcomes.
d. Feed flies and incorporate any desired drugs/treatments until flies reach the desired age for isolation.
B. Preparation of Seahorse calibrant plate and reagents
Note: The calibrant plate and sensor cartridge must be prepared the day prior to the mitochondria isolation to allow 16–24 h for the sensor cartridge to rehydrate.
1. One day before the mitochondria isolation and respiration measurements, prepare the Seahorse calibration plate.
a. Fill each well in the 24-well calibration plate with 1 mL of calibrant solution.
Note: These instructions are written for a Seahorse XF24 analyzer. The calibration plates, cell culture plates, and sensor cartridges contain 24 wells. Volumes should be adjusted if using a 96-well model.
b. Place the sensor cartridge in the filled calibration plate. Make sure the sensors are submerged in the calibrant solution.
c. Store the calibration plate with cartridge in a 37 °C incubator (non-CO2) for 16–24 h.
Note: The sensor cartridges arrive sealed and dry. Complete rehydration of the sensors (for O2 measurements) is essential for proper assay function.
2. Prepare all buffers and injection compound stock solutions the day before the mitochondria isolation and respiration measurements.
C. Mitochondria isolation from whole Drosophila
Note: This section will take approximately 45–60 min to isolate a mitochondria pellet from whole Drosophila. Begin thawing all reagents and stock solutions on ice at this time.
1. Anesthetize flies using CO2.
2. Use a brush to move flies (50–100 per strain/condition) into 1.5 mL Eppendorf tubes.
3. Place tubes on ice to keep flies chilled and unconscious while collecting all flies.
Note: It is not necessary to wash excess cuticle wax prior to the addition of mitochondria isolation buffer.
4. Add 500 μL of ice-cold mitochondria isolation buffer (see Recipes).
5. Homogenize flies using a sterile plastic pestle (Figure 1A, B).
Caution: Perform this step carefully to maintain intact mitochondria. Homogenize using straight up and down motions and gentle pressure. Do not grind the tissue or use a motorized grinder (Figure 1B).
Figure 1. Mitochondria isolation. A–F. Isolation of intact mitochondria by grinding (A) whole flies in (B) isolation buffer and performing (C–F) multiple differential centrifugation steps to clarify a mitochondria pellet.
6. Add an additional 500 μL of ice-cold mitochondria isolation buffer.
7. Centrifuge samples at 300× g for 5 min at 4 °C. This step will pellet any remaining whole flies, limbs, and cuticles (Figure 1C).
Caution: Carefully check centrifugation speeds for steps C7, 9, 11, and 15. The first two centrifugation steps are slow (300× g) to pellet whole cells and debris. The second two centrifugation steps are faster (3000× g) to pellet mitochondria.
8. Transfer supernatant to a clean, labeled 1.5 mL Eppendorf tube. The mitochondria will remain in the supernatant. Take care to avoid transferring any debris.
9. Centrifuge samples at 300× g for 5 min at 4 °C.
10. Transfer supernatant (Figure 1D) to a clean, labeled 1.5 mL Eppendorf tube.
11. Centrifuge the supernatant at 3000× g for 10 min at 4 °C. The mitochondria will be in the pellet after this centrifugation step (Figure 1E).
12. Remove the supernatant.
13. Add 1 mL of mitochondria isolation buffer to the pellet.
14. Resuspend the mitochondrial pellet by gently pipetting up and down with a p1000 pipette.
15. Centrifuge the supernatant at 3,000× g for 10 min at 4 °C.
16. Remove the supernatant. Purified mitochondria are in the pellet (Figure 1F).
17. Resuspend the mitochondria pellet in 100 μL of mitochondria isolation buffer.
a. Resuspend the pellet by gently pipetting up and down.
b. Keep the isolated mitochondria on ice.
Caution: Isolated mitochondria should be used for respiration measurements within 4 h of isolation.
D. Determination of protein concentration in isolated mitochondria
Note: This section will require approximately 30–45 min to complete and calculate results. To increase efficiency, begin preparing standards during centrifugation steps in part C.
1. Create a standard curve using stock BSA diluted in mitochondria isolation buffer. BSA concentrations for the standard curve: 0, 0.2, 0.4, 0.6, 0.8, and 1.0 mg/mL.
2. Dilute isolated mitochondria in isolation buffer. Mitochondria protein concentration must be within the range of the prepared BSA standard curve. Dilutions ranging from 1:10 to 1:100 will typically yield a protein concentration within the linear range of the protein assay.
3. Add 20 μL of Reagent S to 1 mL of Reagent A (both from the Bio-Rad DC Protein Assay kit) to create working Reagent A.
4. Mix the protein assay reagents and diluted standards or mitochondria in a 96-well plate.
a. Set up three separate reaction wells for each standard and mitochondria sample.
b. 5 μL of diluted mitochondria or BSA protein standard.
c. 25 μL of working Reagent A.
d. 200 μL of Reagent B (Bio-Rad DC Protein Assay Kit). Reagent B should be added last. Production of a colored reaction product will begin when Reagent B is added. Work quickly for consistent results.
5. Incubate the protein assay plate at room temperature in the dark for 15 min.
Caution: Color will be stable for up to 1 h after mixing reagents.
6. Measure sample absorbance at 750 nm using a plate reader.
7. Determine protein concentration in mitochondria samples by plotting the BSA standard curve and performing a linear regression.
a. Calculate the mean value for the technical triplicates before performing the linear regression.
b. Make sure to account for dilution of the original mitochondria sample (1:10 or 1:100) when determining the protein concentration of the mitochondria samples.
E. Respiration measurements with isolated mitochondria
Note: Approximately 45 min is required to complete the setup of the Seahorse assay plate, and approximately 60 min is required to complete a Seahorse respiration protocol.
1. Prepare fresh 1× working mitochondria assay buffer with pyruvate and malate.
Note: Substrates other than pyruvate and malate may be used in the assay buffer. See General notes.
2. Dilute mitochondria to 0.1 µg/µL (based on protein concentration assay) in 1× working mitochondria assay buffer (with pyruvate and malate).
3. Obtain a 24-well Seahorse XF24 assay plate.
4. Pipette 50 µL of diluted mitochondria to the bottom of the assay well (5 µg of total protein).
Note: Use technical triplicates for each unique strain/treatment to ensure reproducibility of results.
5. Centrifuge assay plate at 3,000× g for 30 min at 4 °C in a tabletop centrifuge with a swinging microplate rotor for cell culture plates.
6. Dilute injection compound stock solutions in 1× working mitochondria assay buffer to create working injection compounds (Table 1).
Note: Perform these dilutions and load the injection ports of the Seahorse assay cartridge (step E7) while the assay plate is in the centrifuge. The assay cartridge should be ready when the centrifugation step is completed.
a. Perform serial dilutions of injection compound stock solutions to avoid pipetting very small volumes.
b. Injection compound concentrations are calculated based on the desired final concentration in the assay plate. See Table 1 for examples of compound dilutions.
Table 1. Injection compounds for Seahorse analyzer
Injection compound Stock concentration Working concentration Injection volume Final concentration
A. ADP 100 mM 30.66 mM 75 µL 4 mM
B. Oligomycin 10 mM 34.66 µM 75 µL 4 µM
C. FCCP 10 mM 77.32 µM 75 µL 8 µM
D. Rotenone/
Antimycin A
5 mM/10 mM 21 µM/42 µM 75 µL 2 µM/4 µM
Note: See Troubleshooting for suggestions regarding injection compound optimization.
7. Load 75 µL of each injection compound into the injection ports in the assay cartridge (Figure 2A and B).
Figure 2. Layout of Seahorse cartridge injection ports and assay plate. A. Arrangement of the injection ports surrounding a sensor in the Seahorse cartridge for an XF24 analyzer. Each well of the 24-well plate (shown in B) contains a group of four injection ports surrounding the sensor. Each section of the injection port will receive 75 µL of working injection compound. C. Side view of an individual injection port and sensor. D. Plate layout for the 24-well plate model. The wells marked with an X should be filled with assay buffer only, and no mitochondria, to serve as background controls. The background wells are in the default recommended arrangement used in the WAVE software. They may be changed manually, but users should select multiple background wells in different areas of the plate. Created in https://BioRender.com.
8. Use a light microscope to check for mitochondria attachment to the bottom of the Seahorse assay plate. See Figure 3 for an example of a light microscope view of a monolayer of attached mitochondria.
Figure 3. Light microscope image of plated mitochondria after centrifugation of Seahorse assay plate. Panel A and A’ show a confluent monolayer of mitochondria. Panel B and B’ show a plate with a low density of plated mitochondria. A’ and B’ are enlarged images taken from the dashed line boxes in panels A and B.
9. Add 450 µL of 1× working mitochondria assay buffer with pyruvate and malate to each assay well. Pipette carefully to the side of the wells. Do not disturb the mitochondria monolayer on the bottom of the plate.
10. Set up the Seahorse XF24 analyzer respiration experiment.
a. Use the WAVE desktop software to program the experiment.
b. Set the desired assay temperature.
Note: Seahorse analyzer temperatures are dependent on the ambient room temperature of the instrument. The desired experiment temperature should be ~12–20 °C above ambient room temperature. If the desired experiment must be carried out at cooler temperatures, the analyzer may be set up in a cold room.
c. Identify and label sample groups (technical replicates) before starting the instrument protocol.
Note: This step is critical for streamlined data analysis.
d. See Table 2 for an example of the programmed steps for a respiration experiment.
Note: Each step of the respiration measurement should be repeated 2–3 times to demonstrate stable oxygen consumption rates by the isolated mitochondria.
Table 2. Example Seahorse instrument run protocol
Command Time Compound
Calibrate n/a n/a
Equilibrate n/a n/a
Loop 2× n/a
Mix 1 min n/a
Measure 4 min n/a
Mix 30 s n/a
Inject A n/a ADP
Loop 2× n/a
Mix 1 min n/a
Measure 2 min n/a
Mix 30 s n/a
Inject B n/a Oligomycin
Loop 2× n/a
Mix 1 min n/a
Measure 2 min n/a
Mix 30 s n/a
Inject C n/a FCCP
Loop 2× n/a
Mix 1 min n/a
Measure 2 min n/a
Mix 30 s n/a
Inject D n/a Rotenone/Antimycin A
Loop 2× n/a
Mix 1 min n/a
Measure 2 min n/a
Mix 30 s n/a
End
a. First, insert the cartridge and utility plate.
b. Initiate the calibration protocol.
c. After calibration, the Seahorse will eject the utility plate. Remove the utility plate and insert the mitochondria cell culture plate.
Note: Take care to insert the plate in the correct orientation.
d. Once the respiration measurements are complete, discard the assay cartridge and the cell culture plate.
Data analysis
Analysis of the mitochondria protein assay should be performed as quickly as possible while the isolated mitochondria are kept on ice. Linear regression analysis of a standard curve can be performed quickly using Microsoft Excel. Average the absorbance at 750 nm for each standard and sample technical triplicate. Plot the A750 vs. the known protein concentration for the BSA standards in an XY scatter plot in Excel (X = protein concentration, Y = A750). Include a linear regression on the chart and show the formula. Use the calculated formula to determine the concentration of the diluted mitochondria samples. Be sure to account for any dilution of the mitochondria when determining the concentration in the resuspended mitochondria pellet.
Initial analysis of mitochondria respiration is performed automatically by the Seahorse analyzer WAVE Desktop software. Following completion of the instrument run, WAVE will generate data files containing raw data for oxygen consumption rate (OCR), oxygen pressure (O2 mm Hg), and extracellular acidification rate (ECAR), though ECAR is not a useful measurement when assessing isolated mitochondria. The raw data will include a data point for each assay well at every time point measured. Keep these data files for reference. The WAVE software will also automatically average the OCR data for all wells that were grouped as technical replicates. WAVE will also display the average OCR for a group or individual sample well for each measurement cycle. These data may be used to generate data plots in WAVE software. An example of this application is shown in Figure 4. Data from the WAVE Desktop software may be exported to Excel or GraphPad Prism for further analysis.
Figure 4. Sample Seahorse data. Graph of oxygen consumption rates (OCR) of isolated mitochondria plated in a 24-well assay and analyzed using a Seahorse analyzer. Injection compounds are noted by vertical lines. Each point on the graph represents an OCR measurement by the instrument (expressed as pmol oxygen consumption/minute). Each point is an average of three technical replicates for each group, with the exception of the light blue line, which represents the four background wells. The error bars represent the standard error of the mean for the technical replicates. The dark blue line is untreated healthy mitochondria, while the dark red line is mitochondria isolated from cisplatin-treated Drosophila. Statistical analyses were performed using GraphPad Prism. Treatment groups were compared using Welch’s t-test to compare one group directly to another group at each measurement.
The most important analysis of Seahorse data can be performed in Microsoft Excel or GraphPad Prism. Mitochondrial function analysis often uses discreet data points to describe the activity of mitochondria in the presence of inhibitors [9]. State 2 respiration is measured prior to the addition of ADP. This value should be low, as the mitochondria do not have a substrate to generate ATP. The addition of ADP creates state 3 respiration, where mitochondria are actively generating ATP. The addition of oligomycin to block ATP synthase creates state 4o. Mitochondria enter state 3u upon the addition of the uncoupler FCCP. OCR following the final injection of rotenone and antimycin A represents non-mitochondrial respiration (state 4). These respiratory states can then be used to calculate other descriptive measurements of mitochondria respiration (See Table 3).
Table 3. Mitochondria respiration measurement calculations
Measurement Calculation Explanation
Basal respiration State 3–State 4 Normal function of the mitochondria with substrates available.
Maximal respiration State 3u Uncoupled respiration. Maximum OCR for the isolated mitochondria.
Reserve capacity State 3u–State 3 Measure of the additional potential of mitochondria to increase their oxygen consumption.
ATP-linked respiration State 3–State 4o Mitochondria respiration that is generating ATP.
Proton leak State 4o–State 4 Oxygen consumption that does not generate ATP.
Coupling efficiency ATP-linked/basal respiration Estimate of how much of the basal OCR is used to generate ATP.
Respiratory control ratio State 3/State 4o Measure of coupling of OCR to ATP synthesis.
State apparent 4 – (ATP-linked respiration/maximum respiratory capacity) Measure of how close to maximum capacity the mitochondria are.
Calculations of the relevant mitochondria respiration states can be performed in Excel or Prism. Statistical analyses should be performed using software like Prism. The specific tests performed will depend on which samples will be compared. Multiple comparison two-way ANOVA with Holm-Sidak multiple comparisons correction should be used when determining statistical significance between multiple different groups. Unpaired Welch’s t-test may be employed for comparing one strain or treatment to only one other strain. It is useful to perform statistical analyses after calculating the above mitochondria respiration states. Each statistical test will then show if mitochondria from different groups exhibit different respiration at different parts of the experimental protocol. Perform a statistical analysis for each respiration measurement to determine all possible significantly different respiration patterns among the sample groups.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Groen, et al. [16]. Genetic Reduction of Mitochondria Complex I Subunits is Protective Against Cisplatin-Induced Neurotoxicity in Drosophila. Neurobiology of Disease (Figure 6, panel C–K).]
The protocol described here was validated during the completion of experiments for the above publication. In the 2022 article, we used this protocol to demonstrate that different Drosophila strains exhibited different oxygen consumption rates in response to three days of cisplatin treatment, and that improved mitochondria function was correlated with resistance to cisplatin neurotoxicity (see Groen et al. [16], Figure 6C). We validated the isolation of the mitochondria by examining plated mitochondria with a simple light microscope and 20× objective lens to confirm the protocol produced a monolayer in the assay plate (Figure 3). We also produced and fixed a mitochondria pellet, which was prepared and sliced for analysis by transmission electron microscopy. See Figure 5 for representative TEM images confirming that our isolated mitochondria pellets are indeed a highly purified population.
Figure 5. TEM of isolated mitochondria. An isolated mitochondria pellet was fixed and stained for TEM analysis. The pellet was cut after fixation for imaging. Panels A, B, C, D, and E show the isolated mitochondria at 3,000×, 8,000×, 12,000×, 15,000×, and 30,000× magnification, respectively. All images were brightened at 20% to aid visualization.
Validation work for the publication also included testing multiple mitochondria concentrations to identify an optimal quantity of mitochondria that demonstrated a robust response to the injection compounds without completely depleting the oxygen in the sample wells. Once the above validation measures confirmed the procedure yielded an intact, purified population of respiring mitochondria, we completed our analysis of Drosophila strain differences with and without cisplatin treatment. At a minimum, we typically perform technical triplicates within an assay plate and complete each set of experiments a minimum of three times. Example data from one Seahorse XF24 analyzer plate is included in Figure 4. The sample data shows two different treatment conditions (with cisplatin and without cisplatin) with three technical triplicates for each treatment group. The data shows respiring mitochondria with a robust response to each of the four injection compounds and low standard error, demonstrating reproducible results across multiple sample wells. The published figure noted above (Groen et al. [16], Figure 6C) uses results from four separate Seahorse assay plates, each with five replicates for each condition. These data demonstrate the protocol described herein yields reproducible data through multiple replicates performed on different days with independent Seahorse assay plates.
General notes and troubleshooting
General notes
1. This protocol contains detailed step-by-step instructions for one possible method of respiration measurements. The substrates used in the assay buffer (pyruvate and malate) and the injection compounds (oligomycin, antimycin A, rotenone, FCCP, ADP) may be modified or changed depending on the goals of the experiment. We have described a protocol to analyze electron flow in the mitochondria electron transport chain using substrates for Complex I. This analysis used a pyruvate/malate substrate combination. If researchers desire to assess electron flow from Complex II, succinate and rotenone may be used as an initial substrate combination.
2. An Agilent Seahorse XF24 analyzer was used for the experiments described in this protocol. This analyzer model has since been discontinued and replaced by the Seahorse XFe24 model. Assay-specific details, such as the amount of mitochondria, assay buffer volumes, injection compound volumes, and software setup may have to be modified for the instrument that will be used.
3. This protocol provides step-by-step instructions for a Bio-Rad DC protein assay. Users may use any previously validated protein concentration assay for this section.
4. Seahorse injection compounds are selected based on the desired analysis of the electron transport chain in isolated mitochondria. This protocol shows an example of substrates and inhibitors selected to analyze electron flow through the electron transport chain starting with input to complex I. Pyruvate and malate provide substrates for the conversion of acetyl coA and the citric acid cycle to generate electron carriers for the ETC. ADP provides a substrate for ATP synthesis. Oligomycin A stops the ETC and ATP synthesis by blocking ATP synthase activity. FCCP uncouples the ETC from ATP synthesis and allows assessment of maximal respiratory capacity. Rotenone and antimycin A block the activity of complex I and complex III, respectively.
5. Keep all flies, lysates, and mitochondria pellets on ice or in a 4 °C centrifuge throughout the isolation protocol. Only take samples out of ice to complete pipetting/supernatant transfer steps.
Troubleshooting
Problem 1: Mitochondria do not exhibit a change in oxygen consumption rate in response to injection compounds.
Possible cause: Incorrect amount of mitochondria plated, injection compounds are the wrong concentration.
Solution: Try plating multiple amounts of mitochondria and check for a confluent monolayer using a light microscope with a 20× objective. Basal oxygen consumption (before the addition of ADP) should be below 200 pmol/min, as the mitochondria will only be able to use residual ADP in the mitochondria until it is added as an injection compound. Each injection compound concentration will also need to be tested to optimize the OCR response. The suggested range of concentrations to titrate is as follows: ADP (0.5–8 mM), oligomycin (0.5–8 µM), FCCP (2–10 µM), rotenone (0.5–4 µM), and antimycin A (0.5–8 µM).
Problem 2: Unstable OCR measurements. OCR may decline over the 2 min reading for each step of the Seahorse protocol.
Solution: The number of plated mitochondria is likely too high. The oxygen in the assay wells will be used up and cannot replenish fast enough in between mixing steps. Reduce the mitochondria concentration to ensure that the oxygen supply is not exhausted during measurement steps (Figure 6).
Figure 6. Optimization of mitochondria number. Representative Seahorse respiration data for mitochondria isolated from untreated Drosophila. Each line represents a different amount of total mitochondria (determined by protein concentration) plated in the Seahorse assay plate. 50 µg and 25 µg of total protein result in respiration curves that are unstable and rapidly decline during each step, indicating oxygen is consumed too quickly. 10 µg and 5 µg have more stable oxygen consumption rates, but 5 µg shows a more robust response to injection compounds and a smaller standard error of the mean.
Problem 3: High well-to-well variation between technical replicates.
Solution: The mitochondria were likely not mixed well prior to plating. Thoroughly mix the mitochondria in assay buffer by carefully pipetting up and down to completely resuspend and mix the mitochondria.
Problem 4: Mitochondria are not isolated intact.
Solution: Use care when pipetting mitochondria in isolation buffer or assay buffer. Mitochondria should be pipetted slowly with a p1000 pipette to avoid damaging the mitochondria during resuspension steps.
Problem 5: Poor mitochondria function (low OCR readings).
Solution: Perform every step of the isolation on ice or in a 4 C centrifuge. Work quickly to ensure the Seahorse assay is started before the mitochondria population degrades (mitochondria will remain active for 2–4 h after isolation).
Acknowledgments
The authors would like to acknowledge the lab of Dr. Eugenia Trushina at Mayo Clinic for valuable conversations regarding mitochondria respiration analysis. We would also like to thank the Mayo Clinic Microscopy and Cell Analysis Core for preparation and staining of fixed mitochondria pellets for TEM analysis.
Competing interests
The authors declare no competing financial or non-financial interests.
Ethical considerations
All experiments described in this manuscript were performed with Drosophila melanogaster and are not included in any IACUC protocols.
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Received: Oct 5, 2024
Accepted: Nov 28, 2024
Available online: Dec 20, 2024
Published: Feb 5, 2025
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Cell Biology > Cell metabolism > Respirometry
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5,181 | https://bio-protocol.org/en/bpdetail?id=5181&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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Peer-reviewed
Development and Validation of Chlamydia muridarum Mouse Models for Studying Genital Tract Infection Pathogenesis
YW Yihui Wang
ZH Zixuan Han
LW Luying Wang
XS Xin Sun
QT Qi Tian
TZ Tianyuan Zhang
Published: Vol 15, Iss 3, Feb 5, 2025
DOI: 10.21769/BioProtoc.5181 Views: 40
Reviewed by: Andrea GramaticaPooja MukherjeeNidhi MenonJohn P Phelan
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Original Research Article:
The authors used this protocol in Infection and Immunity Jul 2021
Abstract
Animal infection models play significant roles in the study of bacterial pathogenic mechanisms and host–pathogen interactions, as well as in evaluating drug and vaccine efficacies. Chlamydia trachomatis is responsible for infections in various mucosal tissues, including the eyes and urogenital, respiratory, and gastrointestinal tracts. Chronic infections can result in severe consequences such as trachoma-induced blindness, ectopic pregnancy, and infertility. While intravaginal inoculation of C. muridarum mimics the natural route of sexual transmission between individuals, transcervical inoculation allows the organisms to directly infect endometrial epithelial cells without interference from host responses triggered by chlamydial contact or infection of vaginal and cervical cells. Therefore, in this study, we used mouse models to visualize pathologies in both the endometrium and oviduct following C. muridarum inoculation.
Key features
• This protocol develops the mouse-adapted Chlamydia muridarum model, ideal for visualizing pathologies in both the endometrium and oviduct genital tract.
• Requires female mice and utilizes specific techniques for intravaginal and transcervical inoculation with chlamydial elementary body (EB) and a form specialized for intracellular replication.
• The protocol necessitates specialized equipment, including a laminar flow hood, a micropipette, and a non-surgical embryo transfer device (NSET).
Keywords: Chlamydia muridarum Chlamydia trachomatis Mouse model Genital tract infection Pathogenesis
Graphical overview
Background
All chlamydial species are obligate intracellular bacteria that replicate exclusively inside eukaryotic host cells. Chlamydial infections are dependent on a complex infection cycle that depends on transitions between specific forms. This cycle consists of a cell form specialized for host cell invasion, the elementary body (EB), and a form specialized for intracellular replication, the reticulate body (RB). Chlamydia trachomatis is a leading cause of sexually transmitted bacterial infections. Although C. trachomatis is sensitive to antibiotics, many infected individuals neglect medical treatment due to the lack of specific symptoms. The lack of treatment may allow chlamydial organisms to ascend to the upper genital tract, causing pelvic inflammatory diseases, ectopic pregnancy, or tubal infertility, depending on the upper genital tissues affected and the severity of the invasion by the chlamydial organisms. The current clinical challenges include no effective means for preventing chlamydial ascension and no approved C. trachomatis vaccines for preventing chlamydial infection or disease. Modeling chlamydial infection in the female mouse genital tract has provided a productive platform for revealing pathogenic and protective mechanisms during chlamydial infection [1].
The mouse-adapted C. muridarum can efficiently evade innate immunity in the lower genital tract and ascend to the upper genital tract where it induces long-lasting sequelae [2,3] such as hydrosalpinx, which is similar to that observed under laparoscopy in women infected with the human pathogen C. trachomatis. Thus, the C. muridarum model has been used to investigate the pathogenic mechanisms of long-term chlamydia infection [4–8]. In contrast, infection of the female mouse genital tract with C. trachomatis is limited and often fails to induce oviduct pathology [9]. Nevertheless, intravaginal inoculation with a high dose of C. trachomatis or transcervical inoculation with a moderate dose can cause glandular duct dilation, a pathology that was initially identified in C57 mice infected with C. muridarum [10]. Glandular duct dilation is observed as uterine horn dilation macroscopically. Since hydrosalpinx and uterine horn dilation can both lead to infertility, these pathologies have been used for evaluating chlamydial pathogenicity. Therefore, in this study, we used mouse models to study the pathology of the endometrium and oviduct following C. muridarum inoculation.
Materials and reagents
Biological materials
1. Mice: female (15–18 g), ≥ 6 weeks old, with strains and genotypes (wild type, transgenic, or knockout) to be determined depending on the specific experiment and model chosen. Mice were bred and maintained under specific pathogen-free conditions and on a standard chow diet for 8 weeks starting at 6 weeks old in the institutional animal facility of Shanghai Institute of Immunity and Infection
2. HeLa cells (human cervical epithelial carcinoma cells) (ATCC, catalog number: CCL2)
Reagents
1. Medroxyprogesterone acetate (Depo-Provera, Amphastar Pharmaceuticals, Inc, catalog number: 5401); resuspend in sterile 1× PBS (see Recipes) at a final concentration of 25 mg/mL
2. MD-76®R (NDC, catalog number: 0019-1317-09)
3. Trypan blue staining solution (Servicebio, catalog number: G1019-10ML)
4. 0.25% Trypsin-EDTA (1×), (Gibco, catalog number: 25200056)
5. 4% paraformaldehyde (Beyotime, catalog number: P0099-500 ml)
6. 0.1% Triton-X (Biosharp, catalog number: BL934B)
7. 3% BSA (COOLABER SCIENCE&TECHNOLOGY, catalog number: SL1331)
8. Disposable vacuum filters (500 mL, 0.22 μm) (BIOFIL, FMC201500)
9. DMEM (Gibco, catalog number: 11965092)
10. Heat-inactivated fetal bovine serum (FBS) (ExCell, catalog number: FND500)
11. Gentamicin, 10 mg/mL (BBI, catalog number: A428430)
12. Cycloheximide, 10 mg/mL (MCE, catalog number: HY-12320)
13. Sucrose (Sigma-Aldrich, catalog number: S9378)
14. Potassium phosphate, monobasic (Sigma-Aldrich, catalog number: P5655)
15. Potassium phosphate, dibasic trihydrate (Sigma-Aldrich, catalog number: P9666)
16. L-glutamic acid monosodium salt (Sigma-Aldrich, catalog number: G1626)
17. Sodium phosphate, dibasic (BBI, catalog number: A610879)
18. Sodium chloride (BBI, catalog number: A610476)
19. Potassium chloride (BBI, catalog number: A610440)
20. Potassium phosphate, monobasic (BBI, catalog number: A424391)
Solutions
1. Cycloheximide + gentamicin (C&G) growth medium (see Recipes)
2. MD-76R solutions, 52%, 44%, 40% (see Recipes)
3. Sucrose-phosphate-glutamate (SPG) buffer (see Recipes)
4. 1× PBS (see Recipes)
Recipes
1. Cycloheximide + gentamicin (C&G) growth medium
Reagent Volume
DMEM 500 mL
Heat-inactivated fetal bovine serum 50 mL
Gentamicin, 10 mg/mL 1 mL
Cycloheximide, 10 mg/mL 250 μL
Store at 4 °C.
2. MD-76R solutions, 52%, 44%, 40%
Solution Reagents
52% MD-76R 52 mL MD-76R + 48 mL ddH2O
44% MD-76R 44 mL MD-76R + 56 mL ddH2O
40% MD-76R 40 mL MD-76R + 60 mL ddH2O
Adjust to a total volume of 100 mL with sterile water and store at 4 °C.
3. Sucrose-phosphate-glutamate (SPG) buffer
Reagent Quantity
Sucrose 74.62 g
Potassium phosphate, monobasic 0.51 g
Potassium phosphate, dibasic trihydrate 1.23 g
L-glutamic acid monosodium salt 0.82 g
ddH2O 1,000 mL
Total 1,000 mL
Mix with a stir bar until homogenous. Then, filter-sterilize the SPG buffer using disposable vacuum filters (500 mL, 0.22 μm, BIOFIL, FMC201500) and store at room temperature.
4. 1× PBS, pH 7.2
Reagent Quantity
Sodium phosphate, dibasic 1.44 g
Sodium chloride 8 g
Potassium chloride 0.2 g
Potassium phosphate, monobasic 0.24 g
ddH2O 1,000 mL
Total 1,000 mL
Mix with a stir bar until homogenous. Sterilize PBS buffer by autoclaving and store at room temperature.
Laboratory supplies
1. 1 mL syringe & 27-gauge needle (Becton, Dickinson and Company, catalog number: 305109)
2. 1.5 mL microcentrifuge tubes (e.g., Fisher Scientific, catalog number: 21-402-903 or equivalent)
3. Speculum and non-surgical embryo transfer device (NSET) (ParaTechs, catalog number: 60010 or equivalent)
4. 96-well flat bottom cell culture plates, TC-treated (BIOFIL, catalog number: TCP010096)
5. Cell counting plate (Countstar, catalog number: CO010101)
Equipment
1. Microscope (YueHe, model: YHF40)
2. Centrifuge for 96-well plate (Eppendorf, model: Centrifuge 5810 R)
3. Laminar flow hood (Nuaire BSC Class II or equivalent)
4. Multi-channel pipettor (RAININ, model: L12-200XLS+)
5. P20 micropipette (Gilson, model: TB27868 or equivalent)
6. Fluorescence microscope (Olympus, model: AX70) with a CCD camera (Hamamatsu)
7. Ultracentrifuge (Beckman Coulter, model: Optima L-100K) with swinging bucket rotor (Beckman, model: SW 28 Ti or equivalent)
8. Superspeed centrifuge (Thermo Scientific, model: Sorvall RC 6 Plus) with fixed-angle rotor (Sorvall SS 34 or equivalent)
9. Sonics VCX 130 sonicator (Sonics & Materials, model: VCX 130 or equivalent).
10. Countstar Altair (Shanghai Ruiyu Biotechnology Co., Ltd)
Procedure
A. Amplification and purification of chlamydial organisms
1. Chlamydial organisms are amplified in cell culture and purified as EBs using a gradient centrifugation method, as briefly described below:
a. Prepare a gradient from bottom to top in a tube with 5 mL of 52% MD-76R, 8 mL of 44% MD-76R, and 13 mL of 40% MD-76R (see Recipes).
b. Sonicate the infected cells 8–15 times at 60 W for 30 s each to release EBs from cells.
c. Centrifuge the sonicated lysates at 1,200× g for 10 min at 4 °C to remove the cell pellet.
d. Load 10 mL of the supernatant on top of each gradient using the same method as for laying MD-76R layers.
e. Ultracentrifuge at 17,000× g for 90 min at 4 °C in Beckman SW 28 Ti rotor (slow acceleration and brakes off).
f. The EB band can be seen as a cloudy white/yellow band in the 52%–44% interphase after ultracentrifuge.
g. Resuspend gradient-purified EBs in SPG buffer (see Recipes) and store in aliquots at -80 °C until use.
B. Intravaginal inoculation with C. muridarum EBs
1. Five days before inoculation with C. muridarum EBs, clean mouse abdomen with 70% (v/v) ethanol and inject 100 μL of 25 mg/mL medroxyprogesterone acetate with a 27- or 30-gauge needle by subcutaneous injection (I.s). Observe mouse for 2–3 min for any signs of adverse effects.
2. Take a C. muridarum stock aliquot from the -80 °C freezer and thaw on ice. Using sterile techniques, operate the following steps inside a laminar flow hood.
3. Dilute the stock with sterile SPG buffer to the desired 2 × 105 inclusion-forming units (IFUs) per 10 μL. Mix well and keep on ice until mouse inoculation.
Note: For example, if the stock titer is 1 × 107 IFU/µL, first dilute the stock in 1:10 (10 μL + 90 μL SPG buffer) for a stock titer of 1× 106 IFU/µl. Then, continue to dilute the stock in 1:50 (20 μL + 980 μL SPG buffer) for a final stock titer of 2 × 104 IFU/µL (2 × 105 IFUs per 10 μL).
4. Use a p20 micropipette to pipette 10 μL of solution from the tube that contains the desired C. muridarum EBs. Place the micro-pipettor flat on the right-hand side.
5. Use the left hand to hold the mouse’s belly up, and the head tilted downward (Figure 1A).
6. Use the right hand to gently insert the micro-pipettor tip (containing 10 μL of inoculum) into the vagina until a slight resistance is felt (from the cervix). Slowly eject the inoculum until the first stop is reached. Hold the micro-pipettor without releasing the plunger when removing the micropipette tip from the vagina (Figure 1B).
7. Continue to hold the mouse in the same position for 2 min to ensure inoculum remains in the vaginal vault (which may increase chlamydial invasion of vaginal and ectocervical epithelial cells).
8. After the 2 min, the mouse can be returned to the cage. A brief observation for any signs of adverse effects is useful for ensuring the quality of the procedure.
9. For evaluating pathology, euthanize the mice with CO2 for 2 min in the chamber and fix them on the dissection table. Using micro-scissors, incise the abdominal skin, abdominal muscle, and peritoneum along the midline.
10. Lift the intestinal tissue upward to expose the Y-shaped reproductive tract and surrounding adipose tissue. Use micro-scissors to cut the mouse's pubis and intermittently cut the lower section of the reproductive tract below the pubis (Figure 1C).
11. Then, gradually cut and lift the reproductive tract upward, separating it from the surrounding connective tissue.
Figure 1. Key steps of intravaginal inoculation of mice genital tract with C. muridarum. A. Schematic diagram of intravaginal inoculation of C. muridarum in the mice genital tract. Mice were inoculated intravaginally with 2 × 105 IFUs of C. muridarum as described in section A. B. Schematic diagram of local magnification of genital tract infection manipulation in mice. C. Pathology after intravaginal inoculation; a group of mice was inoculated intravaginally with 2 × 105 IFUs of C. muridarum (n = 5, left) or SPG buffer (Ctrl, n = 5, right) as described in section A. On day 56, mice were sacrificed to observe upper genital tract pathology hydrosalpinx. Data from two or three independent experiments.
C. Transcervical inoculation with C. muridarum EBs
1. Five days before inoculation with C. muridarum EBs, clean the mouse’s abdomen with 70% (v/v) ethanol and inject 100 μL of 25 mg/mL medroxyprogesterone acetate intraperitoneally (i.p) with a 27- or 30-gauge needle. Observe the mouse for signs of adverse effects.
2. After five days, take a C. muridarum stock aliquot from the -80 freezer and thaw on ice. Using sterile techniques, operate the following steps inside a laminar flow hood.
3. Dilute stock with sterile SPG buffer to desired 2 × 105 IFUs per 10 μL. Mix well and keep on ice until mouse inoculation.
4. Connect the NSET device to the p20 micropipette and take up 10 μL of chlamydial EBs.
5. Gently place a speculum into the vagina to open up the mouse’s vagina. Insert the NSET device into the speculum and through the cervix. Press the pipette plunger completely to deliver chlamydial EB.
6. Remove the NSET device gently without releasing the pipette plunger and remove the speculum.
7. Monitor mice for several minutes for any adverse signs and then return them to the cage.
8. For evaluating cervix dilation, after euthanizing mice with CO2 in the chamber, fix mice on the dissection table. Using micro-scissors, incise the abdominal skin, abdominal muscle, and peritoneum along the midline.
9. Lift the intestinal tissue upward to expose the Y-shaped reproductive tract and surrounding adipose tissue. Use micro-scissors to cut the mouse's pubis and intermittently cut the lower section of the reproductive tract below the pubis.
10. Gradually cut and lift the reproductive tract upward, separating it from the surrounding connective tissue (Figure 2).
Figure 2. Key steps of transcervical inoculation of mice genital tract with C. muridarum. A. Schematic diagram of transcervical inoculation of C. muridarum in the mice genital tract. Mice were inoculated with 2 × 105 IFUs of C. muridarum as described in section B. B. Schematic diagram of local magnification of transcervical inoculation manipulation in mice. C. Pathology after transcervical inoculation; mice were inoculated with 2 × 105 IFUs of C. muridarum (n = 5, left) or SPG buffer (n = 5, right) as described in section B. On day 56, mice were sacrificed for observing uterine horn dilation. Data from two or three independent experiments.
D. HeLa cells preparation
1. Prepare a 175 cm flask seeded with 1-day-old HeLa cells at approximately 90% confluency.
2. Wash Hela cells once with 1× PBS and treat with 0.25% Trypsin-EDTA (1×) in a CO2 incubator for 2 min. Add 10 mL of DMEM +10% FBS medium to the 175 cm2 flask and mix the cells after Trypsin-EDTA treatment.
3. Set up a HeLa monolayer 96-well microplate with 2.5 × 104 cells/well. Make sure that monolayer confluency is kept between 70%–80% at the time of C. muridarum infection.
Note: 10 μL of cell suspension + 10 μL of trypan blue staining solution (total 20 μL) was automatically counted using a Countstar Altair.
E. Detecting the load of live chlamydial organisms
1. The day after setting up the HeLa monolayer microplate, add 500 μL of sterile SPG buffer and 3–4 autoclaved glass beads to a 1.5 mL microcentrifuge tube in the flow hood.
2. Label the tube with the ID of the mouse that will be swabbed.
3. Insert a sterile rayon mini-tipped applicator into the vaginal vault without pushing past the cervix. A slight resistance can be felt when the swab touches the cervix.
4. Rotate the swab clockwise and counterclockwise five times in each direction.
5. Pull the swab out and place it in the pre-labeled 1.5 mL microcentrifuge tube. Cut the handle of the applicator short enough to close the microcentrifuge cap and immediately put the tube on ice for subsequent processing.
6. Thoroughly vortex swab samples for 2 min at high speed to release chlamydial organisms into solution. Then, prepare a 96-well flat bottom TC-treated plate to be used for serially diluting the swab sample.
7. After serial dilution (from 1:10 to 1:80 by 2-fold serial dilutions), transfer 100 μL of the serially diluted swab samples onto the 96-well plate with 24-hours-old HeLa cell monolayer using a multi-channel pipettor starting from the highest diluted row.
8. Centrifuge the 96-well plates infected with chlamydial samples at 800× g for 1 h at room temperature.
9. Remove inoculum and add 200 μL of C&G growth medium to the 96-well plates.
10. Incubate the plates in a 37 °C, 5% CO2 tissue culture incubator . After 20–24 h of incubation, the microplates are ready for processing to visualize chlamydial inclusions using an immunofluorescence assay.
Note: Here, the primary antibody (rabbit antibody) was made with purified C. muridarum elementary bodies and was a gift from Jingyue Ma from Tianjin Medical University General Hospital. After primary antibody incubation, the secondary antibody (goat anti-rabbit IgG conjugated with Cy2, Abcam, #ab6940) was added, and nuclei were visualized using the DNA dye Hoechst 33258 (Abcam, #ab228550). The doubly labeled samples were used for counting chlamydial inclusions under a fluorescence microscope with a CCD camera.
11. Fix the infected HeLa cells with 100 μL of 4% paraformaldehyde and incubate for 40–60 min at room temperature.
12. Remove paraformaldehyde and wash the plate with 1× PBS once. Then, permeabilize the fixed cell monolayers with 100 μL of 0.1% Triton-X for 30 min at room temperature.
13. Block the permeabilized cells with 100 μL of 3% BSA for 1 h at 37 °C or overnight at 4 °C after removing the Triton-X solution.
14. Label chlamydial antigens with a rabbit polyclonal antibody raised with C. muridarum EBs as the primary antibody: add 50 μL of the rabbit antibody and incubate at 37 °C for 1 h or overnight at 4 °C.
15. Remove the primary antibody solution and wash the plate with 1× PBS three times.
16. Detect the immobilized primary antibody by adding 50 μL of the secondary antibody conjugate (goat anti-rabbit conjugated with FITC) solution that contains Hoechst DNA dye and incubate at 37 °C for 1 h.
17. Remove the secondary antibody solution and wash the plate with 1× PBS three times.
18. Calculate the total number of IFUs per swab or tissue based on the number of inclusions per well, dilution factor, and inoculation volumes (see sects E12–16).
F. Live chlamydial stock titration
1. One day prior to titration, seed cells at 1×105 cells per well in a 24-well tissue culture plate at approximately 90% confluency.
2. Make sure that monolayer confluency is kept between 70%–80% at the time of live chlamydial stock titration.
3. Prepare a 1:100 stock dilution by adding 10μL of EB stock to 990μL of SPG buffer. The starting 1:10,000 dilution was prepared by adding 10μL of 1:100 stock dilution to 990μL of SPG buffer. A 10-fold serial dilution was set up for as many serial dilutions as required (Figure 3A).
4. After serial dilution (from 104 to 109, by 10-fold serial dilutions), transfer 200 μL of the serially diluted samples onto the 24-well plate with 24-hour-old HeLa cell monolayer (Figure 3B).
5. Centrifuge the 24-well plates infected with chlamydial samples at 800× g for 1 h at room temperature.
6. Remove inoculum and add 200 μL of C&G growth medium to the 24-well plates.
10. Incubate the plates in a 37 °C, 5% CO2 tissue culture incubator for 20–24 h.
11. The microplates are ready for processing to visualize chlamydial inclusions using an immunofluorescence assay (see steps D10–17) (Figure 3C, D).
12. Count inclusions under a fluorescence microscope as follows.
13. Starting with the highest dilution at 10× magnification, count the number of inclusions in the center, upper-right, lower-right, lower-left, and upper-left sections of the well.
14. If fewer than 10 inclusions are visible per field at 10× magnification, count the inclusions present in three wells. Conversely, if more than 25 inclusions are visible, use the next highest magnification and count inclusions.
15. Evaluate each dilution at the lowest magnification, increasing the magnification accordingly. Count five random views and record the dilution and magnification for each count.
16. Determine IFU/mL as described below:
IFU/mL=(IFU counted/Fields counted) (Dilution)[ πr2/π(A/2)2]/Inoculation vol (mL)−1
Where: r2=(well radius)2
FOV=field of vision
A=FOV eye piece/objective magnification
π(A/2)2=Area of radius viewed
Note 1. When sample infection rate reaches 100%, these samples are not included in the count.
Note 2. The cell monolayer was not used to count the number of inclusions (does not cover the entire field or has swaths missing).
Note 3. If your sample is diluted appropriately, an inclusion is the result of an EB infection of a HeLa cell, which is designated as an inclusion-forming unit or IFU.
Figure 3. Live chlamydial titration by immunofluorescence assay. A. Prepare the chlamydia stock. B. HeLa cells are infected by several dilutions for 22–24 h. C. The cell plate is set up by immunofluorescence assay. D. The fluorescence signal is detected under a fluorescence microscope.
Data analysis
The evaluation of hydrosalpinx in mice following intravaginal inoculation is performed as follows (Table 1).
Table 1. Hydrosalpinx score
Score Pathology
0 No hydrosalpinx
1 Hydrosalpinx only observable under magnification
2 Visible hydrosalpinx smaller than the size of ovary
3 Visible hydrosalpinx roughly equal the size of ovary
4 Visible hydrosalpinx larger than the size of ovary
Validation of protocol
This protocol was validated by the Guangming Zhong lab, and the resulting work was published in (Figures 4, 5):
Sun et al. [10]. Chlamydia muridarum induction of glandular duct dilatation in mice. Infect Immun.
Chen et al. [3]. Chlamydial induction of hydrosalpinx in 11 strains of mice reveals multiple host mechanisms for preventing upper genital tract pathology. PLoS One.
Figure 4. Data example showing hydrosalpinx development in 11 strains of mice following lower genital tract infection with C. muridarum. Representative image from each strain of mice is presented with the whole genital tract in the left and the magnified oviduct/ovary portion in the right. The number of mice with positive hydrosalpinx (as marked with red arrows in the magnified oviduct/ovary images) were counted and recorded as a percentage of hydrosalpinx-positive mice. Reprinted/adapted from Chen et al. [3]. 2014 The Authors. Published by PLoS One.
Figure 5. Data example showing C. muridarum induction of uterine horn dilation. Reprinted/adapted from Sun et al. [10]. 2015 The Authors. Published by Infection and Immunity.
General notes and troubleshooting
1. For the gradient purification of EBs, all solutions and centrifuge tubes must be pre-chilled at 4 °C or on ice prior to usage; they should remain on ice during usage. Rotors and buckets utilized during centrifugation must be cleaned with 70% ethanol and pre-chilled prior to use.
2. All steps must be performed aseptically in biological safety cabinets to avoid contamination.
3. For the intravaginal inoculation, suggested inoculation concentrations are 2 × 105 for C. muridarum. It is recommended to titrate your stock in mice before formal experiments.
4. Ensure the micropipette tip is not inserted past the cervix during intravaginal inoculation as this can cause unnecessary pain and injury to the mouse.
5. For the transcervical inoculation, follow the manufacturer’s instructions when using the NSET device.
Acknowledgments
We would like to express our heartfelt thanks to Prof. Guangming Zhong for his invaluable guidance and support throughout our research. We also wish to acknowledge the contributions of his lab members, particularly Jianlin Chen, Hongbo Zhang, Zhou Zhou, and Zhangsheng Yang, for their assistance and collaboration in improving the techniques used in this study. Their expertise and dedication have been instrumental in the success of our work. This work was supported by the National Natural Science Foundation of China for the Youth (32100162) to Q.T. and (32000138) to T.Z.
Competing interests
We declare no conflict of interest or competing interests.
Ethical considerations
All animal studies have been approved by the Ethics Committee of the Institute of Immunity and Infection of Shanghai Chinese Academy of Sciences (A2020019).
References
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Campbell, J., Huang, Y., Liu, Y., Schenken, R., Arulanandam, B. and Zhong, G. (2014). Bioluminescence Imaging of Chlamydia muridarum Ascending Infection in Mice. PLoS One. 9(7): e101634. https://doi.org/10.1371/journal.pone.0101634
Chen, J., Zhang, H., Zhou, Z., Yang, Z., Ding, Y., Zhou, Z., Zhong, E., Arulanandam, B., Baseman, J., Zhong, G., et al. (2014). Chlamydial Induction of Hydrosalpinx in 11 Strains of Mice Reveals Multiple Host Mechanisms for Preventing Upper Genital Tract Pathology. PLoS One. 9(4): e95076. https://doi.org/10.1371/journal.pone.0095076
Tian, Q., Zhou, Z., Wang, L., Abu-Khdeir, A. M., Huo, Z., Sun, X., Zhang, N., Schenken, R., Wang, Y., Xue, M., et al. (2020). Gastrointestinal Coinfection Promotes Chlamydial Pathogenicity in the Genital Tract. Infect Immun. 88(4): e00905–19. https://doi.org/10.1128/iai.00905-19
Conrad, T. A., Gong, S., Yang, Z., Matulich, P., Keck, J., Beltrami, N., Chen, C., Zhou, Z., Dai, J., Zhong, G., et al. (2016). The Chromosome-Encoded Hypothetical Protein TC0668 Is an Upper Genital Tract Pathogenicity Factor of Chlamydia muridarum. Infect Immun. 84(2): 467–479. https://doi.org/10.1128/iai.01171-15
Yang, C., Lei, L., Collins, J. W. M., Briones, M., Ma, L., Sturdevant, G. L., Su, H., Kashyap, A. K., Dorward, D., Bock, K. W., et al. (2021). Chlamydia evasion of neutrophil host defense results in NLRP3 dependent myeloid-mediated sterile inflammation through the purinergic P2X7 receptor. Nat Commun. 12(1): 5454. https://doi.org/10.1038/s41467-021-25749-3
Yang, C., Kari, L., Lei, L., Carlson, J. H., Ma, L., Couch, C. E., Whitmire, W. M., Bock, K., Moore, I., Bonner, C., et al. (2020). Chlamydia trachomatis Plasmid Gene Protein 3 Is Essential for the Establishment of Persistent Infection and Associated Immunopathology. mBio. 11(4): e01902–20. https://doi.org/10.1128/mbio.01902-20
Dockterman, J., Reitano, J. R., Everitt, J. I., Wallace, G. D., Hendrix, M., Taylor, G. A. and Coers, J. (2024). Irgm proteins attenuate inflammatory disease in mouse models of genital Chlamydia infection. mBio. 15(4): e00303–24. https://doi.org/10.1128/mbio.00303-24
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Article Information
Publication history
Received: Aug 3, 2024
Accepted: Nov 28, 2024
Available online: Dec 26, 2024
Published: Feb 5, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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5,182 | https://bio-protocol.org/en/bpdetail?id=5182&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
A Bio-protocol resource
Peer-reviewed
Optimal Dual RNA-Seq Mapping for Accurate Pathogen Detection in Complex Eukaryotic Hosts
IM Infanta Saleth Teresa Eden M.
UV Umashankar Vetrivel
Published: Vol 15, Iss 3, Feb 5, 2025
DOI: 10.21769/BioProtoc.5182 Views: 234
Reviewed by: Prashanth N SuravajhalaSoumya MoonjelyWilly R Carrasquel-Ursulaez
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Abstract
Dual RNA-Seq technology has significantly advanced the study of biological interactions between two organisms by allowing parallel transcriptomic analysis. Existing analysis methods employ various combinations of open-source bioinformatics tools to process dual RNA-Seq data. Upon reviewing these methods, we intend to explore crucial criteria for selecting standard tools and methods, especially focusing on critical steps such as trimming and mapping reads to the reference genome. In order to validate the different combinatorial approaches, we performed benchmarking using top-ranking tools and a publicly available dual RNA-Seq Sequence Read Archive (SRA) dataset. An important observation while evaluating the mapping approach is that when the adapter trimmed reads are first mapped to the pathogen genome, more reads align to the pathogen genome than the unmapped reads derived from the traditional host-first mapping approach. This mapping method prevents the misalignment of pathogen reads to the host genome due to their shorter length. In this way, the pathogenic read information found at lesser proportions in a complex eukaryotic dataset is precisely obtained. This protocol presents a comprehensive comparison of these possible approaches, resulting in a robust unified standard methodology.
Key features
• Benchmarking of top-ranking software for quality control, adapter trimming, and read mapping.
• Emphasizes the importance of read mapping criteria for dual RNA-Seq datasets: (i) high count of uniquely host mapped reads, (ii) low count of host multi-mapped reads, and (iii) high count of unmapped reads belonging to pathogens.
• Elaborates the best mapping approach to precisely extract the pathogen reads as these get captured comparatively less in dual RNA-Seq datasets.
Keywords: Dual RNA-Seq Host–pathogen interactions Host-first mapping Pathogen-first mapping Misalignment of pathogen reads
Graphical overview
Background
Dual RNA sequencing is a powerful tool to precisely investigate the complete gene expression profile of two actively interacting organisms. When a pathogen infects a host, both develop adaptation mechanisms by revealing a series of changes at molecular levels. These biological changes are interrelated and can be effectively elucidated by analyzing the whole transcriptome data of the host–pathogen interaction. Until 2012, transcriptomics sequencing of infectious diseases was limited to separately capturing the cascade of biological events in either the host or the pathogen [1]. This allows us to study gene expression changes only from a linear perspective. Over the years, insightful research in transcriptomics analysis has encouraged researchers across the globe to devise the concept of extracting and sequencing whole RNA from the infection site. This idea will provide a broader scope for unraveling the role of smaller molecules and studying the biological changes in organisms at different stages of interaction. Compared to conventional RNA sequencing technology, dual RNA sequencing caters to a sufficient amount of host and pathogen RNA and other small RNA molecules (microRNA, long non-coding RNA, and other non-coding RNA) when pathogen-infected samples are subjected to the well-established and published methods of RNA extraction and sequencing procedures [2–5].
Dual sequencing data generated as reads or fragments comprises information on both the host and the pathogen. The infected host cells may not always reflect pathogen-related data in enormous quantities. Therefore, a detail-oriented analysis is necessary to capture the minimally found essential information on host–pathogen interplay. Several bioinformatics analysis methods and tools are available to explore the complex datasets involving eukaryotic and prokaryotic reads [6–8]. Most of these methods suggest a sequential mapping approach to extract host and pathogen reads. Some studies recommend a combined mapping approach where the reference genomes of the host and pathogen are concatenated, indexed, and used as a single reference [9,10]. As far as quality control and adapter removal are concerned, standard methods for read mapping have been receiving greater attention as pathogen data is found in lesser quantities. Espindula et al. [10] conducted studies on different infection models and showed that other alternative mapping approaches outperform the traditional host-first mapping approach. When trimmed reads are mapped first to the host reference, there are high chances of pathogen read mismapping due to its shorter read length. To avoid this, alternative mapping approaches were introduced, and this bio-protocol emphasizes one such mapping technique—the pathogen-first mapping approach—in detail. A comparison of mapping results proved that most of the pathogen reads have been restored in the pathogen-first mapping approach. The results obtained using this approach are now based on a higher confidence scale and can be used for further processing.
This protocol is presented with the aim of demonstrating a standardized bioinformatics procedure for productive mapping. In this protocol, human monocyte-derived macrophages (HMDMs) infected with Mycobacterium tuberculosis (Mtb) were used as a dual RNA-Seq model. Apart from humans infected with Mtb, other host and pathogen models can still be used in dual RNA-Seq experiments. For example, other dual RNA-Seq datasets are publicly available in Gene Expression Omnibus (GEO); one such dataset is Triticum aestivum infected with Fusarium graminearum (Accession: SRP439529; GEO: GSE233409). Dual RNA-Seq test dataset of human–Mtb was downloaded from Sequence Read Archive (SRA), National Centre for Biotechnology Information (Accession: SRP359986 [11]). This study primarily focuses on a compound that restricts Mycobacterium tuberculosis from catabolizing cholesterol by binding with iron. The data quality control before and after read trimming was performed using FastQC. The FastQC tool can be accessed online at https://www.bioinformatics.babraham.ac.uk/projects/fastqc/. For trimming low-quality bases and adapter removal, benchmarking was performed using the topmost adapter trimming software, namely fastp [12] and Trim-Galore (https://github.com/FelixKrueger/TrimGalore). Following this, the topmost splice-aware alignment tools STAR [13] and HISAT2 [14] were also benchmarked to obtain optimal results. The most important part of the analysis is to apply the best mapping method that serves the experimental purpose of dual RNA-Seq analysis. After mapping the reads to their respective genomes, the reads mapping to genes were quantified using featureCounts [15]. Other downstream analysis methods, which include read count normalization, differential expression analysis, gene ontology, and pathway enrichment, are not demonstrated here, as the key aim of this protocol is to highlight the crucial preparatory steps like quality control, adapter removal, and read mapping in a descriptive manner.
Software and datasets
1. Data
Dual RNA-Seq datasets are publicly available on NCBI Sequence Read Archive (SRA) and can be downloaded and analyzed for learning purposes. For this protocol, a dataset from SRA (accession: SRP359986, Gene Expression Omnibus datasets; GEO: GSE196816) was downloaded and utilized.
2. Bioinformatics tools (all tools were installed using conda)
• SRA-Toolkit (version 3.1.0) includes tools like prefetch and fasterq-dump for fetching SRA datasets and extracting individual fastq files.
• FastQC (version 0.12.1). For assessing the quality of fastq reads before and after trimming.
• MultiQC (version 1.19). For integrating results into interactive visualization reports throughout the analysis.
• TrimGalore (version 0.6.10) and Cutadapt (version 4.6). For quality-trimming bases from reads, automatic adapter detection and removal, and filtering reads based on lengths.
• HISAT2 (version 2.2.1). For indexing the reference genome and mapping trimmed high-quality reads to the reference genome of eukaryotes.
• BWA (version 0.7.17-r1188). For indexing the reference genome and mapping trimmed high-quality reads to the reference genome of prokaryotes.
• SAMtools (version 1.19). For converting huge files from mapping results (.sam) into binary formatted .bam files enabling easy processing, to sort reads with their mate pairs, and to check the statistical distribution of reads after mapping.
• Bedtools (version v2.31.1). For extracting the interleaved reads inside .bam files into paired-end separate fastq files.
• featureCounts from Subread package (version v2.0.6). For quantifying all reads mapped to genomic coordinates using annotation feature file(.gtf/.gff3) of the reference genome.
3. Platform used: Linux, Ubuntu
• CPU: Architecture, 64 bit; 24 cores, 96 threads
• Memory: 512 GB RAM
Note: The threads/cores mentioned in each step of the analysis need to be modified by users as per the computational resources available.
Procedure
Pseudocode for the steps used in the analysis:
START OF ANALYSIS
# Step 1: Downloading SRA Datasets and Preparation of raw fastq files
DOWNLOAD the list of SRA accession ids
CREATE a file with SRA accession list
USE ‘prefetch’ command on the SRA list to download data files
USE ‘fasterq-dump’ command to extract fastq files from the downloaded data files
USE ‘gzip’ command to compress extracted fastq files in .fastq.gz format
# Step 2: Initial Quality Control of raw fastq files
USE ‘fastqc’ command to check quality of raw data
USE ‘multiqc’ to create consolidated QC report of raw data
# Step 3: Data cleaning and Final Quality Control of trimmed data:
USE ‘trimgalore’ command to trim adapter reads and low-quality bases from raw reads
USE ‘fastqc’ command to check quality of adapter-trimmed reads
USE ‘multiqc’ command to create consolidated QC report of adapter-trimmed reads
# Step 4: Mapping high-quality fastq reads to the reference genome:
CREATE the hg38 reference genome index using ‘hisat2’
CREATE the Mtb reference genome index using ‘bwa’
USE ‘bwa’ to map the adapter-trimmed reads to Mtb genome index
USE ‘samtools’ to extract the unmapped reads from the generated .bam files
USE ‘bedtools bamtofastq’ to convert .bam files to .fastq files
USE ‘hisat2’ to map the unmapped reads to hg38 genome index
# Step 5: Quantification of reads mapped to genomic features:
USE ‘featurecounts’ on the mapped .bam files to count reads belonging to transcripts/genes/exons/
# Step 6: Downstream transcriptome analysis:
USE the readcounts table to perform gene expression analysis using statistical methods like DESeq2, edgeR or Cufflinks-Cuffdiff
USE tools and databases like ‘BINGO’, ‘CytoHubba’, ‘DAVID’, to identify ontologies and pathways of differentially expressed genes
USE homology search tools like ‘BLAST’ to annotate the differentially expressed genes
END OF ANALYSIS
A. Downloading SRA datasets and preparation of raw fastq files
The sequence datasets used were obtained from NCBI SRA from a dual RNA sequencing study conducted by Theriault et al. [11], where they identified a compound that restricts Mtb from catabolizing cholesterol by binding with iron. In the study, HMDMs were infected with Mtb, and the infected cells were exposed to the following drug treatments: 5 μg/mL ethambutol, 67.5 ng/mL isoniazid, 10 μM mCLB073, 10 μM sAEL057, and dimethyl sulfoxide (DMSO; untreated). For demonstrative purposes, we chose to work with DMSO (untreated) for the control group, and 10 μM mCLB073 and 10 μM sAEL057 for the treatment groups (Table 1). The dataset included both paired-end and single-end read samples; hence, the protocol demonstrates processing both types of libraries in each step.
Table 1 lists the SRR IDs, library type, and treatment given for all samples. All nine sample runs were downloaded from SRA using the SRR run IDs.
Table 1. Sample information
SRR run IDs Library Treatment (Mtb-infected human macrophages)
SRR18042662 Paired DMSO; untreated control
SRR18042663 Single DMSO; untreated control
SRR18042664 Single DMSO; untreated control
SRR18042665 Paired 10 μM mCLB073
SRR18042666 Single 10 μM mCLB073
SRR18042667 Single 10 μM mCLB073
SRR18042668 Paired 10 μM sAEL057
SRR18042669 Single 10 μM sAEL057
SRR18042670 Single 10 μM sAEL057
The sequence reads data were downloaded using the prefetch tool from SRA-Toolkit (version 3.1.0). SRA-Toolkit can be downloaded from https://trace.ncbi.nlm.nih.gov/Traces/sra/sra.cgi?view=software. The paired-end and single-end reads were extracted from the downloads using fasterq-dump from SRA-Toolkit.
Code snippet for downloading RNA-Seq datasets from NCBI-SRA:
$ prefetch SRR18042662 SRR18042663 SRR18042664 SRR18042665 SRR18042666 SRR18042667 SRR18042668 SRR18042669 SRR18042670
$ fasterq-dump --threads 50 SRR18042662 SRR18042663 SRR18042664 SRR18042665 SRR18042666 SRR18042667 SRR18042668 SRR18042669 SRR18042670
Subsequently, the paired and single-end fastq files extracted from SRA downloads were compressed to .fastq.gz format using gzip. The compressed fastq files are the raw reads that will be utilized for the downstream analysis.
B. Initial quality control using FastQC
The quality of reads and bases from the raw FASTQ files were assessed using FastQC. This tool checks the number of reads and their quality, the number of bases and their quality, the presence of adapters, and other statistics such as read length distribution and GC content. FastQC generates separate visualization reports for forward and reverse-read files in HTML format. The HTML files from all samples are then consolidated into a single interactive visualization report using MultiQC [16]. Figures 1 and 2 represent the basic statistics and adapter content before trimming of SRR18042662 sample dataset using FastQC.
Figure 1. Initial QC: Basic statistics for sample SRR18042662
Figure 2. Initial QC: Adapter content before trimming for the sample SRR18042662
Code snippet for performing initial quality check on raw Fastq read files:
$ fastqc -t 10 *.fastq.gz
$ multiqc .
where t is the number of threads to be used to run FastQC. The “.” in the multiqc command represents the current directory.
C. Data cleaning and final quality check
The sequence reads in the raw .fastq.gz files generally exhibited good quality. However, some low-quality bases and adapters are present in the sample reads, which need to be trimmed from the 3' end. After quality control and adapter trimming, the length of the reads varied widely. This variation may cause some reads to lose their mate pairs. Therefore, in addition to trimming, it is also essential to filter reads based on length (default cutoffs: TrimGalore - 20; fastp - 15).
Trimming tools such as fastp (version 0.23.4) and TrimGalore can automatically detect and trim adapters. We used both tools to determine the best results. After trimming, we checked the quality of reads using FastQC. Figures 3 and 4 represent the basic statistics and adapter content of SRR18042662 sample dataset after trimming.
Figure 3. Final QC: Basic statistics for the sample SRR18042662 after adapter trimming
Figure 4. Final QC: Adapter content after trimming for sample SRR18042662
Code snippet for trimming adapters using TrimGalore and Fastp:
# A. TrimGalore
# For paired-end reads
$ trim_galore --quality 20 --stringency 7 --paired SRR18042662_1.fastq.gz SRR18042662_2.fastq.gz --output_dir 2.Trimmed_reads/ --cores 2 --retain_unpaired
# For single-end reads
$ trim_galore --quality 20 --stringency 7 SRR18042663.fastq.gz --output_dir 2.Trimmed_reads/ --cores 2
# B. Fastp
# For paired-end reads
$ fastp -i SRR18042662_1.fastq.gz -I SRR18042662_2.fastq.gz -o SRR18042662_1_clean.fastq.gz -O SRR18042662_2_clean.fastq.gz --detect_adapter_for_pe --cut_tail --correction --overrepresentation_analysis --json SRR18042662.fastp.json --html SRR18042662.fastp.html --thread 16
# For single-end reads
$ fastp -i SRR18042663.fastq.gz -o SRR18042663_clean.fastq.gz --cut_tail --overrepresentation_analysis --json SRR18042663.fastp.json --html SRR18042663.fastp.html --thread 16
TrimGalore parameters:
--quality 20: Trim bases from the ends of reads based on low Phred Score quality (< 20).
--stringency 7: Trims adapter sequences from ends only if there is an overlap of 7 or more bases with the adapter sequence.
--paired: parameter for specifying paired-end reads as input.
--retain unpaired: Removal of low-quality bases and adapters will lead to some reads having very low read lengths. These reads fail to meet length cutoffs and are removed. Their mate pairs remain as single reads and are retained in separate files.
Fastp parameters:
--detect_adapter_for_pe: Automatic adapter detection for paired-end sequences. If disabled, adapter detection will be assumed for single-end sequences.
--cut_tail: trims read at 3' end based on low base quality scores by moving a sliding window from 5' to 3'.
--correction: For paired-end data; read pairs are overlapped to find proper matches, and bases with low quality on one read are corrected to a high quality of their corresponding base on the other read.
--overrepresentation analysis: gives information on where the detected overrepresented sequences are mostly distributed.
When comparing the results obtained from TrimGalore and fastp, we observed that TrimGalore retained almost 99.5% of reads across all samples. Although the Q20 and Q30 Phred values improved with fastp, more reads and bases were removed after trimming (see Table 2 below). Therefore, we chose to proceed with TrimGalore as the preferred trimming tool. We then executed FastQC and MultiQC on the trimmed reads to ensure these were of high quality for downstream analysis.
Table 2. Read-base distribution before and after trimming using Fastp and TrimGalore
Sample Raw Reads Raw read bases (GB) Fastp-trimmed reads Fastp-trimmed bases (GB) TrimGalore-trimmed reads TrimGalore-trimmed bases (GB)
SRR18042662 92,046,490 13.807 88,159,790 (95.78) 12.861 91,532,448 (99.44) 13.376
SRR18042663 28,312,422 2.407 26,720,747 (94.38) 2.194 26,932,291 (95.13) 2.213
SRR18042664 31,636,161 2.689 31,455,656 (99.43) 2.669 31,612,738 (99.93) 2.680
SRR18042665 200,335,050 30.050 192,054,572 (95.87) 27.590 199,221,410 (99.44) 28.677
SRR18042666 33,538,680 2.851 33,242,144 (99.12) 2.821 33,120,616 (98.75) 2.791
SRR18042667 35,517,838 3.019 35,303,710 (99.40) 2.996 35,487,051 (99.91) 3.007
SRR18042668 121,942,300 18.291 115,871,486 (95.02) 16.789 120,703,024 (98.98) 17.522
SRR18042669 30,961,231 2.632 30,760,768 (99.35) 2.610 30,834,234 (99.59) 2.594
SRR18042670 49,600,226 4.216 49,297,844 (99.39) 4.183 49,569,155 (99.94) 4.202
Note: In the above table, the numbers represented in parenthesis indicate the percentage of reads retained after trimming with fastp and trimgalore.
D. Mapping high-quality reads to reference genomes
The next step is to map the trimmed reads to a reference genome (Human - hg38) to identify the genomic locations of all the reads. For genome mapping, there are two leading splice-aware alignment tools: STAR and HISAT2. STAR is a super-fast, highly accurate, and memory-intensive splice-aware aligner. STAR reports a high proportion of uniquely mapped reads as compared to any other topmost splice-aware alignment tools. STAR also maps the non-contiguous reads to the reference genome by a technique called soft clipping. This method improves mapping accuracy, and almost every read is mapped to the genome. However, most of the reads tend to multimap at different genomic locations. There is a chance that the shorter pathogenic reads from the dual RNA-Seq dataset will be multi-mapped to the host genome, and these may also be reported as uniquely mapped reads by soft clipping. In order to avoid such mismapping of reads, we emphasize certain critical checkpoints that are discussed in detail in the following sections of the protocol.
The following criteria are defined for mapping dual RNA-Seq data:
• A high number of uniquely mapped reads.
• A low number of multi-mapping reads.
• A higher number of unmapped reads (likely belonging to a pathogen).
Although both STAR and HISAT2 yield good results independently, we validated the above criteria by mapping our data with both tools.
1. When comparing RefSeq and Ensembl annotations, we chose to work with RefSeq, as it covers most genes with its simplest version of genome annotation [17]. Therefore, as the next step, we downloaded the reference genome assembly (hg38) and annotation file (.gtf) from NCBI RefSeq - GCF_000001405.40. Go to the link to download the reference genome of Homo sapiens: https://www.ncbi.nlm.nih.gov/datasets/genome/GCF_000001405.40/
2. In the above link, click the download button. Then, from the popup screen, select RefSeq only, and choose to download it.
3. Uncompress the downloaded file and collect the reference genome (.fna) and annotation files (.gtf and .gff3). Store these files in a separate folder.
Code snippet for indexing reference genome using STAR and HISAT2:
# Indexing Reference Genome using STAR
$ STAR --runThreadN 4 --runMode genomeGenerate --genomeDir STAR_index --genomeFastaFiles GCF_000001405.40_GRCh38.p14_genomic.fna --sjdbGTFfile genomic.gtf
# Indexing Reference Genome using HISAT2
# Extracting Splice Sites
$ extract_splice_sites.py genomic.gtf > splice_sites.tsv
# Extracting Exons
$ extract_exons.py genomic.gtf > exons.tsv
# Building Genome Index using exon and splice site information
$ hisat2-build GCF_000001405.40_GRCh38.p14_genomic.fna hisat2_hg38_index/hg38_index -p 40 --ss splice_sites.tsv --exon exons.tsv
STAR indexing parameters:
--runThreadN 4: Number of cores to use for processing. It is best to use a minimum number of threads/cores, either 4 or 2, as STAR runs faster, exhausting more RAM.
HISAT2 indexing parameters:
--extract_splice_sites.py: To extract and incorporate splice site information from .gtf so that transcripts do not map mostly to intronic regions and also for identifying transcript isoforms.
--extract_exons.py: To extract exon sites from .gtf, so that transcripts map exactly to exonic regions on the genome.
Code snippet for reference genome mapping using STAR and HISAT2:
# Mapping to the Reference Genome using STAR
# Paired-end reads
$ STAR --genomeDir STAR_index --runThreadN 2 --readFilesCommand zcat --readFilesIn SRR18042662_trimmed_1.fq.gz SRR18042662_trimmed_.fq.gz –outFileNamePrefix B.Mapping_STAR/SRR18042662__STAR --outSAMtype BAM Unsorted
# Single-end reads
$ STAR --genomeDir STAR_index --runThreadN 2 --readFilesCommand zcat --readFilesIn SRR18042663_trimmed.fq.gz --outFileNamePrefix B.Mapping_STAR/SRR18042663__STAR --outSAMtype BAM Unsorted
# Mapping to the Reference Genome using HISAT2
# Paired-end reads
$ hisat2 --dta -x hisat2_hg38_index/hg38_index -1 SRR18042662_trimmed_1.fq.gz -2 SRR18042662_trimmed_2.fq.gz -p20 --un-conc-gz B.Mapping_HISAT2/SRR18042662_Unmapped_pairs --summary-file B.Mapping_HISAT2/SRR18042662_align_stats.txt | samtools view -@15 -b -S | samtools sort -n -@15 -o B.Mapping_HISAT2/SRR18042662.sorted.bam -O BAM
# Single-end reads
$ hisat2 --dta -x hisat2_hg38_index/hg38_index -U SRR18042663_trimmed.fq.gz -p20 --un-conc-gz B.Mapping_HISAT2/SRR18042663_Unmapped_reads.fq.gz --summary-file B.Mapping_HISAT2/SRR18042663_align_stats.txt | samtools view -@15 -b -S | samtools sort -n -@15 -o B.Mapping_HISAT2/SRR18042663.sorted.bam -O BAM
The --dta parameter in HISAT2 reports the alignments as the transcript assemblers used to provide. After mapping all our samples with both STAR and HISAT2, the mapping results were tabulated in Table 3 for comparative analysis of the mapping performances.
Table 3. Comparative mapping analysis of HISAT2 vs. STAR
HISAT2 vs. STAR* Total reads Unique Multi-mapped (should be lesser) Unmapped (should be more for dual RNA-Seq data)
SRR18042662 45766224 31950488 (69.81%) 5220616 (11.41%) 8595120 (18.78%)
34890020 (76.24%)* 6217589 (13.59%)* 4658615 (10.18%)*
SRR18042663 26932291 15352634 (57.00%) 6347595 (23.57%) 5232062 (19.43%)
15427880 (57.28%)* 6761705 (25.1%)* 4742706 (17.61%)*
SRR18042664 31612738 21878890 (69.21%) 5409698 (17.11%) 4324150 (13.68%)
22095054 (69.89%)* 5731876 (18.13%)* 3785808 (11.98%)*
SRR18042665 99610705 39819023 (39.97%) 41065084 (41.23%) 18726598 (18.80%)
38181484 (38.33%)* 52467481 (52.67%)* 8961740 (8.99%)*
SRR18042666 33120616 21182314 (63.96%) 7473519 (22.56%) 4464783 (13.48%)
21391317 (64.59%)* 7870514 (23.76%)* 3858785 (11.65%)*
SRR18042667 35487051 24954036 (70.32%) 5706459 (16.08%) 4826556 (13.60%)
25238589 (71.12%)* 6073050 (17.11%)* 4175412 (11.77%)*
SRR18042668 60351512 37169324 (61.59%) 11599955 (19.22%) 11582233 (19.19%)
40350621 (66.86%)* 14124605 (23.4%)* 5876286 (9.74%)*
SRR18042669 30834234 20287934 (65.80%) 6285903 (20.39%) 4260397 (13.82%)
20528529 (66.58%)* 6817201 (22.11%)* 3488504 (11.31%)*
SRR18042670 49569155 37725432 (76.11%) 8155703 (16.45%) 3688020 (7.44%)
38124908 (76.91%)* 8671102 (17.5%)* 2773145 (5.6%)*
Note: In the above table, * refers to the STAR tool, the corresponding number of reads, and percentage of reads mapped using STAR.
From Table 3, it is evident that although STAR produced more uniquely mapped reads in most samples, it is noteworthy that HISAT2 generated fewer multi-mapped reads and more unmapped reads across all samples compared to STAR. Therefore, we can use HISAT2 for mapping reads to eukaryotic genomes.
Assuming the unmapped reads belong to pathogens, we mapped these to the Mtb genome (RefSeq - GCF_000195955.2) using the BWA aligner [18] to obtain the mapping percentages shown below in Table 4 (second column).
Another alternative mapping approach is to map the adapter-trimmed reads directly to the Mtb genome first. This approach is justified because prokaryotic reads are fewer and shorter compared to eukaryotic reads. When using the previously described mapping method, there is a risk of missing some of the prokaryotic reads, as these shorter reads may misalign with the eukaryotic genome. Therefore, mapping to the Mtb genome first will increase confidence in our mapping steps (Table 4). Subsequently, we extracted the unmapped reads and aligned them to the hg38 genome using HISAT2 (Table 5).
This method was followed earlier by Espindula et al. [10], and significant changes were observed in the number of reads mapped using all possible approaches. This protocol extensively demonstrates the same pathogen-first mapping method by elaborating the steps in a more detailed manner with all the code snippets, which can be altered and used.
Code snippet for alternative pathogen-first mapping approach:
# Indexing Mtb genome using BWA short-read aligner
$ bwa index GCF_000195955.2_ASM19595v2_genomic.fna
# Alternative Mapping Approach
# A. Mapping trimmed reads first to Mtb Genome using BWA.
# Paired-end reads
$ bwa mem Mtb_genome/GCF_000195955.2_ASM19595v2_genomic.fna SRR18042662_trimmed_1.fq.gz SRR18042662_trimmed_2.fq.gz -t 20 | samtools view -@15 -b -S | samtools sort -n -@15 -o B.Mapping_BWA_PathogenFirst/SRR18042662.sorted.bam -O BAM
# Single-end reads
$ bwa mem Mtb_genome/GCF_000195955.2_ASM19595v2_genomic.fna SRR18042663_trimmed.fq.gz -t 20 | samtools view -@15 -b -S | samtools sort -n -@15 -o B.Mapping_BWA_PathogenFirst/SRR18042663.sorted.bam -O BAM
# B. Extract unmapped reads using samtools
$ samtools view -b -f 4 B.Mapping_BWA_PathogenFirst/SRR18042662.sorted.bam > B.Mapping_BWA_PathogenFirst/SRR18042662_unmapped.sorted.bam
# C. Convert sorted bam files to fastq files (paired and single-end)
$ bedtools bamtofastq -i SRR18042662_unmapped.sorted.bam -fq SRR18042662_unmapped_Host_1.fq -fq2 SRR18042662_unmapped_Host_2.fq
$ samtools fastq SRR18042663_unmapped.sorted.bam > SRR18042662_unmapped_Host.fq
# D. Map the unmapped (Host-reads) to hg38 using HISAT2
# Refer HISAT2 mapping command (paired and single-end) from previous snippets.
This way of mapping the reads will result in appropriately aligned prokaryotic reads so that we can more accurately use them to study gene expression levels of both the pathogen and the host.
Table 4. Comparison of mapping between pathogen-first and unmapped Mtb reads (from HISAT2)
Sample No. of unmapped reads (pathogen) from HISAT2 results mapped to Mtb genome No. of adapter-trimmed reads mapped to Mtb (pathogen-first method)
SRR18042662 (PE) 4448104 (25.88%) 4715080 (5.15%)
SRR18042663 3859948 (73.77%) 3859968 (14.33%)
SRR18042664 2251961 (52.08%) 2252019 (7.12%)
SRR18042665 (PE) 10709925 (28.60%) 12181384 (6.11%)
SRR18042666 1831236 (41.02%) 1831276 (5.53%)
SRR18042667 2433725 (50.42%) 2433770 (6.86%)
SRR18042668 (PE) 6380085 (27.54%) 6883877 (5.70%)
SRR18042669 1514060 (35.54%) 1514089 (4.91%)
SRR18042670 1507197 (40.87%) 1507299 (3.04%)
From Table 4, we can visualize the significant increase of ~0.1–0.5 million in mapping numbers in the case of paired-end reads, whereas for single-end reads, there is an increase of around 100 reads. Some of the reads may not map in pairs in the case of paired-end reads. Hence, the mapping percentages are lower in paired-end reads compared to single-end reads. The mapping percentages are even lower in the pathogen-first mapping approach since we are using the complete host–pathogen trimmed reads directly for mapping to the Mtb genome.
Table 5. Comparison of mapping between host-first and unmapped host reads (from BWA)
Sample No. of reads mapped to hg38 (host-first approach) No. of unmapped reads (host) mapped to hg38 from BWA results (pathogen-first approach)
Unique Multi-mapped Unmapped Unique Multi-mapped Unmapped
SRR18042662 31950488 (69.81%) 5220616 (11.41%) 8595120 (18.78%) 31893407 (69.68%) 5144240 (11.24%) 6320738 (13.81%)
SRR18042663 15352634 (57.00%) 6347595 (23.57%) 5232062 (19.43%) 15352703 (57.00%) 6347634 (23.56%) 1371986 (5.09%)
SRR18042664 21878890 (69.21%) 5409698 (17.11%) 4324150 (13.68%) 21879065 (69.20%) 5409463 (17.11%) 2072191 (6.55%)
SRR18042665 39819023 (39.97%) 41065084 (41.23%) 18726598 (18.80%) 39652369 (39.80%) 40503121 (40.66%) 13305107 (13.35%)
SRR18042666 21182314 (63.96%) 7473519 (22.56%) 4464783 (13.48%) 21182318 (63.95%) 7473434 (22.56%) 2633588 (7.95%)
SRR18042667 24954036 (70.32%) 5706459 (16.08%) 4826556 (13.60%) 24954231 (70.31%) 5706225 (16.07%) 2392825 (6.74%)
SRR18042668 37169324 (61.59%) 11599955 (19.22%) 11582233 (19.19%) 37076766 (61.43%) 11444024 (18.96%) 8361031 (13.85%)
SRR18042669 20287934 (65.80%) 6285903 (20.39%) 4260397 (13.82%) 20287963 (65.79%) 6285867 (20.38%) 2746315 (8.90%)
SRR18042670 37725432 (76.11%) 8155703 (16.45%) 3688020 (7.44%) 37725559 (76.10%) 8155417 (16.45%) 2180880 (4.39%)
In Table 5, we compare the mapping results of unmapped reads extracted from the BWA results with HISAT2 mapping results from Table 3. We observe that the unmapped reads from the pathogen-first approach have a good number of uniquely mapped reads and is marginally increased for single-end reads. The number of multi-mapped reads is also marginally lower compared to the host-first mapping approach. Unmapped reads are minimal, as we initially mapped the reads to the Mtb genome. Though a significant increase/decrease in the number of host reads may not be observed during comparison, the host reads from the pathogen-first mapping approach are still found to have impactful results while performing downstream analysis.
Therefore, we consider approach B (pathogen-first mapping) as the best approach and HISAT2 for mapping to the eukaryotic genome when working with Dual RNA-Seq data.
Note: There may still be some reads left unmapped after mapping to genomes of both the host and the pathogen. There may even be a possibility of occurrence of horizontal gene transfer (HGT) events, wherein, other bacteria or viruses other than the pathogen of interest may have invaded the host cells. In order to check this, users may extract only the unmapped reads and map these reads against genomes of other species using BLAST search.
E. Read count quantification using featureCounts
In the previous section, we mapped all the reads to their respective genomes. The next step is to count the number of reads to determine which gene or exonic region each read has mapped to the genome. Therefore, we performed a quantification step for both the pathogen-mapped and host-mapped reads using the RefSeq annotation files (.gtf) of both Mtb and hg38. The featureCounts tool was used to count reads that are mapped to exonic regions and genes.
Code snippet for quantification of read counts:
# Read Count Quantification: FeatureCounts - Pathogen read counting
# Paired-end reads
$ featureCounts -a Mtb_Genome/genomic.gff -t 'gene' -g 'Name' SRR18042662.sorted.bam SRR18042665.sorted.bam SRR18042668.sorted.bam -p -o C.Read_Count_Quantification/Pathogen_Quant_PE_Togene_genename.tsv -O --countReadPairs -T 40
# Single-end reads
$ featureCounts -a Mtb_Genome/genomic.gff -t 'gene' -g 'Name' SRR18042663.sorted.bam SRR18042664.sorted.bam SRR18042666.sorted.bam SRR18042667.sorted.bam SRR18042669.sorted.bam SRR18042670.sorted.bam -o C.Read_Count_Quantification/Pathogen_Quant_SE_Togene_genename.tsv -O -T 40
# Merge .tsv files from both paired-end and single-end quantification results.
$ cut -f1,7- Pathogen_Quant_PE_Togene_genename.tsv | grep -v "#" > temp_Pathogen_Quant_PE.tsv
$ cut -f1,7- Pathogen_Quant_SE_Togene_genename.tsv | grep -v "#" > temp_Pathogen_Quant_SE.tsv
$ join -o auto -e '0' -a 1 -a 2 -1 1 -2 1 temp_Pathogen_Quant_PE.tsv temp_Pathogen_Quant_SE.tsv > Pathogen_Quantification_matrix.tsv
# Read Count Quantification: FeatureCounts - Host read counting
# For paired-end and single-end reads (Host), follow the above code snippets by replacing "genomic.gff" file with RefSeq hg38 annotation file (.gff3)
featureCounts parameters:
-t ‘gene’: reads will be mapped to the “gene” feature from the annotation file. Default: ‘exon’.
-g ‘Name’: the quantified reads will be grouped under gene names from .gff if “Name” is mentioned. Default: ‘gene_id’.
-p: To specify that the reads are paired-end
-O: A parameter for quantifying reads with minimum overlapping bases also.
--countReadPairs: By mentioning this parameter, fragments will be quantified instead of reads. This is applicable for paired-end reads.
Join command parameters:
-o auto: A parameter to apply a simple format to the output file.
-e ‘0’: Parameter to fill empty values with zero.
-a: Parameter to fill the file numbers.
-1 1: Join files based on column 1 for file 1.
-2 1: Join files based on column 1 for file 2.
The read count results obtained from the previous step can be used to infer the genes that are differentially expressed under different treatment conditions.
Validation of protocol
The number of reads retained after each step in the analysis significantly impacts data quality, especially when dealing with a dataset containing reads from multiple species. In this protocol, we have identified and listed the top-performing software predominantly used for such analyses. Additionally, we have tabulated the results obtained at each step after thoroughly benchmarking these software tools. Besides the software chosen for trimming and mapping in this protocol, the other sections were validated in earlier studies, which include mapping strategy [10] and choice of genome annotation [17].
General notes and troubleshooting
General notes
1. In this protocol, we have detailed the quality control, mapping, and quantification of read counts specific to both host and pathogen, as these are the most crucial steps in generating highly confident data for downstream analysis. The read count results from these steps are obtained after proper validation at each stage of analysis.
2. We are not demonstrating further downstream analysis in detail here, as it would be a repetition of Bio-protocol references [19–22], and the statistical preferences vary widely based on individual research perspectives.
3. The mapping strategies discussed are of vital importance. By using a pathogen-first approach, we ensure higher confidence in mapping steps, particularly by reducing misalignment issues with shorter prokaryotic reads. This strategy also helps in accurately identifying pathogen reads prior to mapping the remaining reads to the host genome using HISAT2. Such meticulous mapping strategies are critical for obtaining reliable data for efficient downstream analyses.
4. The raw read counts obtained from quantification can be further normalized using the DESeq2 package in R, which applies the median of ratios method of normalization. This method accounts for sequencing depth and RNA composition without considering gene length, as differential expression (DE) analysis compares read counts between sample groups for the same gene. The calculation of median ratios and the script used to obtain normalized counts are also available online at https://hbctraining.github.io/DGE_workshop/lessons/02_DGE_count_normalization.html.
5. DESeq2 works with sample replicates and tests how the variances calculated from read count data of replicates are dispersed. There are several methods for estimating the dispersion based on the nature of the dataset. Generally, in case of fewer replicates, the data may get adjusted to fit even the outliers into the dispersion model; this in turn affects the downstream analysis and biases the gene expression levels. Therefore, it is advisable to have more replicate samples so that the accuracy of data is maintained and the outliers are easily removed. DESeq2 uses negative binomial distribution by default, which is the most widely used dispersion method, as it is extensively designed for biological systems exhibiting excessive variability. Whenever there are unequal numbers of replicates, generalized linear models (GLMs) can be used, as they are flexible to model unequal variances.
6. In the case of the dataset having non-replicate samples, edgeR can be used to perform differential expression analysis:
Code snippet for DEG analysis using edgeR:
# Install and load edgeR library and count databases
install.packages(“edgeR”)
library(edgeR)
readcount <- read.table(“count_data.tsv”, header=TRUE, row.names=1)
# Create DEG object
dds <- DGEList(counts=readcount, group=c(“control”,”treatment”))
# The above line creates a DEG object using the readcounts of one control and one treatment sample.
# Data normalization
dds <- calcNormFactors(dds, method=”TMM”)
# Data normalization using Trimmed means of M-values method.
# Estimation of Dispersion
dds <- estimateDisp(dds, robust=TRUE)
# Fitting negative binomial model
fit <- glmFit(dds, design=model.matrix(~group))
# DEG analysis
deg <- glmLRT(fit, coef=2)
# Extracting significantly expressed genes based on p-value cutoffs
deg_results <- deg[deg$table$PValue < 0.05, ]
7. After normalization, differential gene (DE) expression analysis can be performed by comparing treatment and control groups from the sample dataset. The DE analysis results can be validated by appropriate statistical tests (Wald test, DESeq2), with significantly expressed genes marked by p-values, false discovery rate (FDR), and log-fold changes. The steps for performing DE analysis are described in the Bio-protocol by Hoerth et al. [23] and Fernández et al. [24].
8. After analyzing gene expression levels across several conditions, significantly upregulated and downregulated genes can be functionally annotated using homology search by BLAST.
9. Gene ontology analysis can be performed using BiNGO, a Cytoscape-based plugin, discussed in detail by Duarte et al. [22]. The same method can be followed to predict the functional role of differentially expressed genes. As an alternative, CytoHubba, another cytoscape plugin, can also be used for this purpose. This tool ranks the topmost hub genes out of the significantly expressed genes.
10. Pathway enrichment of differentially expressed genes can also be performed using online tools like DAVID, as mentioned by Chemello et al. [20]. Graphite Web [25] is another resource for pathway enrichment analysis.
Troubleshooting
1. The software and datasets section of the manuscript represents the corresponding latest version numbers, dated as of while performing this analysis. Therefore, it is always recommended that you install and use the latest version of the software with all the bug fixes.
2. While installing software using conda, there are possibilities for already existing versions of the software to be upgraded/downgraded automatically in order to manage package dependencies. Some tools like trimgalore require FastQC and cutadapt to be installed separately, as version compatibility conflicts could arise. In such cases, users can still try to switch the order of software installation to have the latest version in usage.
3. After trimming, it is essential to check whether low-quality bases and adapters are completely removed from the raw reads. The FastQC reports after trimming need to be checked for residual adapter content that may be present. In some cases, the poly-A tail and other bases can still be found. In this case, raw data can be again subjected to trimming using trimgalore by refining the parameters for adapter trimming.
4. While extracting the unmapped reads from .bam files, it is important to ensure that the reads are properly fetched as paired-end, without missing out any reads because of the absence of mapping pairs. In order to address this, the .bam files used for extracting the reads need to be properly sorted by “name” by using “samtools sort -n“ and not by “coordinates”.
GitHub page links for the above-mentioned software:
1. TrimGalore: https://github.com/FelixKrueger/TrimGalore
2. SAMtools: https://github.com/samtools/samtools?tab=readme-ov-file
Acknowledgments
We thank Theriault et al. [11] for sharing the dual RNA-Seq datasets of human monocyte-derived macrophages infected with Mycobacterium tuberculosis at NCBI SRA. We are also thankful to Indian Council of Medical Research (ICMR) intramural Biomarker study grant (NIRT/Intra/5/2023/CD dated 31.01.2024) under which this protocol was demonstrated.
Competing interests
There are no conflicts of interest or competing interest.
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Article Information
Publication history
Received: Aug 28, 2024
Accepted: Dec 8, 2024
Available online: Dec 26, 2024
Published: Feb 5, 2025
Copyright
© 2025 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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Abstract
Recent advancements in high-throughput functional genomics have substantially enhanced our comprehension of the genetic and molecular dimensions of cancer, facilitating the identification of novel therapeutic targets. One of the key methodological innovations in this field is the CRISPR screening strategy, which has proven efficacy in elucidating essential gene functions and pathway alterations critical to cancer cell survival and fitness. The construction of custom CRISPR libraries permits the integration of tailored single-guide RNAs (gRNAs), offering greater flexibility as well as specificity in comparison to the commercially available libraries, and enables more refined secondary screening strategies to attenuate the selection of false positive potential gene candidates. Among various molecular cloning techniques, circular polymerase extension cloning (CPEC) has emerged as a highly efficient and cost-effective approach. CPEC utilizes polymerase overlap extension to assemble overlapping DNA fragments into circular plasmids, eliminating the need for restriction digestion and ligation and thus streamlining the creation of both single and multi-fragment constructs. In this protocol, we present the application of the CPEC method to construct the EpiTransNuc knockout gRNA library, specifically designed to target epigenetic regulators, transcription factors, and nuclear proteins. The custom library, assembled using the lentiGuide-Puro backbone, comprises 40,820 gRNAs, with 10 gRNAs per gene, along with 100 non-targeting control gRNAs. Importantly, the CPEC method can be tailored to meet the specific requirements of other custom gRNA libraries, offering flexibility for diverse research applications.
Key features
• Involves PCR-based linearization of the backbone with designed primer sets.
• Facilitates flexibility in gRNA composition and number in library construction.
• Skips conventional cloning techniques such as restriction digestion and ligation.
Keywords: Circular polymerase extension cloning (CPEC) CRISPR libraries CRISPR screening circular DNA assembly guide RNAs (gRNAs)
Graphical overview
Schematic representation of the circular polymerase extension cloning (CPEC) procedure
Background
Our knowledge of the genetic and molecular aspects of cancer has advanced considerably owing to high-throughput functional genomics, which facilitates the identification of potential treatment targets or therapeutic vulnerabilities for a range of cancer types [1]. Genome-wide or focused CRISPR screening strategy has proven to be a versatile tool to unravel genes and even alterations in pathways having a major role in the survival and fitness of cancer cells, as well as therapy resistance [2,3]. Researchers can uncover details about gene functions, interactions, and regulatory networks by creating libraries containing different CRISPR constructs [4]. This understanding is crucial for deciphering complex biological systems and diseases.
Despite the availability of genome-wide or focused CRISPR libraries from commercial or repository sources, the creation of custom-built CRISPR libraries holds paramount importance for several reasons. First and foremost, de novo library construction enables the incorporation of a specific number of gRNAs, rather than being constrained by a predetermined or limited quantity. In contrast to genome-wide commercial libraries, which typically include a limited number of gRNAs per gene, custom-built libraries allow for a higher number of gRNAs per gene, thereby improving the reliability of the observed phenotypes. Moreover, to refine results and eliminate false negative candidates, secondary screens that focus on potential hits identified in primary screens are often conducted. To accomplish this, constructing a de novo library using gene sets that have already been pre-screened would become increasingly important. Second, the composition of the library can be adjusted with respect to the focus of the research. Although this study explores the CRISPR library within the context of cancer research, its application extends beyond this field. To exemplify, engineering gRNAs targeting transcription factors, metabolic genes, and immune regulation genes can be prioritized based on the specific research focus. Last but not least, a vector backbone with a Cas9 expression cassette or one that lacks Cas9 can be chosen.
In this context, researchers exploit cloning methodologies, which are classified into two primary categories: those that are sequence-dependent (Gateway cloning) and those that are sequence-independent [5], typically based on homologous recombination. Sequence-dependent strategies utilize restriction digestion-ligation methods or site-specific recombination and necessitate unique and specific sites within the insert, vector, or both. This dependence on unique sites makes these approaches less ideal for multi-fragment cloning. On the other hand, sequence-independent approaches comprise of ligation independent cloning (LIC) [6], sequence and ligation-independent cloning (SLIC) [7], Gibson assembly [8], and covalently-closed-circular synthesized (3Cs) [9]. All of these methods have pros and cons, including circular polymerization extension cloning (CPEC); however, CPEC could be exploited for the construction of CRISPR libraries whenever someone opts for a more cost-effective and streamlined approach.
Circular polymerase extension cloning (CPEC) represents a relatively cost-effective, high-efficiency cloning technique compared to the Gibson assembly and other protocols used in molecular cloning. It operates on the principle of polymerase overlap extension and is considered a robust alternative due to its omission of restriction digestion, ligation, and other procedural steps [10]. CPEC enables the transformation of overlapping DNA fragments into a double-stranded circular form via the polymerase extension mechanism, thereby facilitating the integration of the insert into the target plasmid. During the CPEC reaction, linear double-stranded inserts and vectors are first separated through increasing temperature (denaturation). Subsequently, the resulting single-stranded products anneal through their overlapping regions and use each other as templates to construct the circular plasmid. Through the CPEC method, not only a single gene but also multi-fragment assembly could be obtained. From this point of view, CPEC could have a pivotal role in the construction of CRISPR libraries (de novo). In this method, it is crucial to uniquely select the overlapping regions of the insert and vector, and the melting temperature (Tm) should be as high as possible to minimize the likelihood of vector self-ligation and concatenation.
Here, we present a robust, cost-effective, and straightforward CPEC-based protocol to engineer a custom-built gRNA library for CRISPR screening research. The knockout gRNA library, enriched for transcription factors and epigenetic regulators as well as nuclear factors, hence named EpiTransNuc, was constructed by exploiting the PCR linearized lentiGuide-Puro backbone and comprises 40820 gRNAs, comprising 10 gRNAs per gene and 100 non-targeting controls.
Materials and reagents
Biological materials
1. Endura electrocompetent E. coli bacteria (Lucigen, catalog number: 60242-1)
Reagents
1. lentiGuide-Puro backbone (Addgene, catalog number: 52963)
2. Designed primer sets for lentiGuide-Puro backbone linearization:
a. Forward primer: GTTTTAGAGCTAGAAATAGCAAGTTAAAATAAGGCTAGTCCGTTATCAACTTGAAAAAGTGGCACC
b. Reverse primer: CGGTGTTTCGTCCTTTCCACAAGATATATAAAGCCAAGAAATCGAAATACTTTCAAGTTACGG
3. Q5 reaction buffer (New England Biolabs, catalog number: B9027S)
4. Q5 high GC enhancer (New England Biolabs, catalog number: B9028A)
5. 10 mM dNTPs (New England Biolabs, catalog number: N0447S)
6. Q5 high-fidelity DNA polymerase enzyme (New England Biolabs, catalog number: M0491S)
7. Ultra-pure DNase/RNase-free distilled water (Invitrogen, catalog number: 10-977-015)
8. NucleoSpin gel and PCR clean-up (Macherey-Nagel, catalog number: 740609.50)
9. EpiTransNuc library synthesized by CustomArray (Genscript) (the gRNA pool was improved upon the Addgene #51047 nuclear proteins gRNA sub-pool library through the inclusion of gRNAs from Sabatini, Brunello, GecKO, and Toronto v3)
10. Absolute ethanol (Isolab, catalog number: 9.200.262.500)
11. 10× FastDigest Esp3I (BsmBI, IIs class) enzyme (Thermo Fisher Scientific, catalog number: FD0454)
12. 10× FastDigest buffer (Thermo Fisher Scientific, catalog number: FD0454)
13. DTT (Neofroxx, catalog number: 1114GR005)
14. NEBNext high-fidelity PCR master mix (New England Biolabs, catalog number: M0541S)
15. Agarose Biomax (Prona, catalog number: 000320PR)
16. Gel loading dye, purple 6× (New England Biolabs, catalog number: B7024SVIAL)
17. 1 kb plus DNA ladder (New England Biolabs, catalog number: N3200L)
18. Isopropanol (Isolab, catalog number: 961.023.2500)
19. Electroporation cuvettes (Bio-Rad, catalog number:1652089)
20. Tryptone (AppliChem, catalog number: 1553.03)
21. Yeast extract (Nzytech, catalog number: MB16401)
22. NaCl (Merck, catalog number: M106404.1000)
23. LB broth with agar (Sigma-Aldrich, catalog number: L2897)
24. Ampicillin (Sigma-Aldrich, catalog number: A0166)
25. Tris base (Sigma-Aldrich, catalog number: T1503-1KG)
26. EDTA (Sigma-Aldrich, catalog number: E5134-500G)
27. Glacial acetic acid (Sigma-Aldrich, catalog number: 27225-2.5L-R)
28. SafeView Classic (ABM, catalog number: G108)
29. Endotoxin-free HiSpeed Plasmid DNA Maxi kit (Qiagen, catalog number: 12362)
Solutions
1. 10× Tris-acetate-EDTA buffer stock solution (10× TAE) (see Recipes)
2. Luria Bertani (LB) agar stock solution (see Recipes)
3. Luria Bertani (LB) liquid medium stock solution (see Recipes)
Recipes
Note 1: For the preparation of recipes, in-house ddH2O was used.
Note 2: A 10× TAE solution is used as stock and is stable at room temperature. Prepare LB agar stock and LB liquid medium stock solutions right before usage and keep them at 4 °C.
1. 10× Tris-acetate-EDTA buffer stock solution (10× TAE)
Reagent Final concentration Quantity or Volume
Tris base 400 mM 48.4 g
EDTA 9.9 mM 3.7 g
Glacial acetic acid 1.14% (v/v) 11.4 mL
ddH2O - 800 mL
Total n/a Complete with ddH2O to 1,000 mL
2. LB agar stock solution
Reagent Final concentration Quantity or Volume
LB broth with agar 3.5% (w/v) 35 g
ddH2O - 1,000 mL
Total n/a 1,000 mL
3. LB liquid medium stock solution
Reagent Final concentration Quantity or Volume
Tryptone 1% (w/v) 10 g
Yeast extract 0.5% (w/v) 5 g
NaCl 1% (w/v) 10 g
ddH2O 1,000 mL
Total n/a 1,000 mL
Laboratory supplies
1. Nunc square bioassay dishes (Thermo Fisher Scientific, catalog number: 240835)
2. 0.2 mL PCR tubes (Labselect, catalog number: PT-02-C)
3. 1.5 mL Eppendorf (Golden Gate, catalog number: KG2211)
4. Petri dish Ø90 × 17 mm (Isolab, catalog number: 081.02.191)
5. Drigalski glass spreader (Superior, catalog number: C180024)
Equipment
1. SimpliAmp thermal cycler (Applied Biosystems, model: A24811)
2. NanoDrop2000 spectrophotometer (Thermo Fisher Scientific, model: ND-2000)
3. Microcentrifuge (Thermo Fisher Scientific, model: MicroCL 17R 230 V)
4. Balance (Sartorious, model: ENTRIS 822-1S)
5. Micropulser electroporator (Bio-Rad, catalog number: 1652100)
6. Gel electrophoresis system (Thermo Fisher Scientific, model: OWL B1A)
7. Gel Doc XR+ system with Image Lab software (Bio-Rad, model: 1708195)
8. 37 °C bacterial orbital shaker (Thermo Fisher Scientific, model: 4329)
9. 37 °C incubator (Heraeus, model: Heracell)
10. Heat block (Thermo Fisher Scientific, model: Digital Shaking Drybath)
11. Centrifuge (Eppendorf, model: 5810R)
Software and datasets
1. Python v2.7.17
2. Biopython v1.79
3. Count_spacers.py code (code is deposited at https://github.com/fengzhanglab/Screening_Protocols_manuscript)
4. BioRender (https://www.biorender.com/). The following figures were created using BioRender: Graphical overview, Senturk, S. (2025) https://BioRender.com/o79n165
Procedure
Circular polymerization extension cloning (CPEC) protocol comprises four major steps:
A. Preparation of lentiGuide-Puro backbone (Addgene #52963) via PCR-based linearization.
B. Preparation of gRNA library (in our research, every gene is targeted by 10 gRNAs).
C. Bulk cloning of amplified gRNA library into lentiGuide-Puro backbone by pursuing CPEC methodology.
D. Transformation of the gRNA cloned lentiGuide-Puro backbone into electrocompetent Endura bacteria.
A. Preparation of the lentiGuide-Puro backbone via PCR-based linearization
1. The primary objective of this methodology is to eliminate all stages that might compromise product purity and, by extension, the cloning process.
Critical: Based on our experimental assessments, gel extraction is identified as the critical initial step that needs to be addressed. From this regard, the major technical aim is to minimize or circumvent gel purification steps wherever technically feasible. To facilitate this, the lentiGuide-Puro backbone was initially linearized through PCR-based linearization.
Note: Linearize 1 ng of lentiGuide-Puro backbone with the primer pairs shared below (Table 1).
Table 1. Primer pair for backbone linearization
Primer pair Sequence
Forward primer GTTTTAGAGCTAGAAATAGCAAGTTAAAATAAGGCTAGTCCGTTATCAACTTGAAAAAGTGGCACC
Reverse primer CGGTGTTTCGTCCTTTCCACAAGATATATAAAGCCAAGAAATCGAAATACTTTCAAGTTACGG
2. Mix gently the reaction components as well as primer pairs designed specifically for the linearization of lentiGuide-Puro and incubate for 24 cycles following the conditions detailed below. Perform the linearization step with Q5 polymerase enzyme on ice by using 0.2 mL PCR tubes.
Note: Prepare a master mix using a 1.5 mL Eppendorf tube and then divide the reaction into 0.2 mL PCR tubes.
3. Reaction components comprise 10 μL of reaction buffer (Q5), 10 μL of GC enhancer buffer, 1 μL of dNTP (10 mM stock), 3 μL of forward primer, 3 μL of reverse primer (10 μM stock), 0.5 μL of Q5 polymerase enzyme, a maximum amount of 1 ng of lentiGuide-Puro plasmid, and ddH2O adding up to the total volume of 50 μL (Table 2).
Table 2. PCR-based backbone linearization conditions
Reagents Final concentration Volume required (μL)
5× Q5 reaction buffer 1× 10
5× Q5 HighGC enhancer 1× 10
10 mM dNTP 200 μM 1
10 μM forward primer 0.6 μM 3
10 μM reverse primer 0.6 μM 3
Q5 high-fidelity DNA polymerase 0.02 U/μL 0.5
Lenti-guidePuro backbone Maximum 1 ng
ddH2O Varies
Total 50
4. Follow the thermocycling conditions shown in Table 3.
Table 3. Thermocycling conditions for the PCR reaction
Step Temp. (°C) Duration No. of cycles
Initial denaturation 98 3 min 1
Denaturation 98 10 s
24
Annealing 60.9 30 s
Extension 72 8 min
Final extension 72 10 min 1
Hold 4 ∞ -
5. Perform in total 10 different PCR linearization reactions.
6. Pool those 10 reactions and column purify samples using NucleoSpin Gel and PCR clean-up kit.
7. To detail this section, first add ethanol into concentrated buffer NT3. Then, mix previously pooled and purified samples and buffer NT1 supplied by the kit in a 1:2 ratio. Insert NucleoSpin Gel and PCR clean-up column into the collection tubes (2 mL) supplied with the kit and then load a 700 μL sample. Perform all steps and centrifugation at room temperature. Centrifuge samples at 11,000× g for 30 s. This step is followed by the removal of the flowthrough. Wash silica membrane with 700 μL of buffer NT3 twice and subsequently centrifuge samples at 11,000× g for 30 s. After removal of the flowthrough, dry silica membranes by centrifugation again at 11,000× g for 1 min.
Note 1: Since residual ethanol may interfere with downstream enzymatic reactions, columns were incubated at 70 °C. In the elution step, transfer columns into new 1.5 mL Eppendorf tubes and keep samples initially with 15–30 μL of buffer NE (composition of elution buffer NE: 5 mM Tris/HCl, pH 8.5) for at least 1 min, followed by centrifugation at 11,000× g for 1 min).
Note 2: The product resulting from the linearization PCR measures 8,303 bp, and its termini align with the Esp3I (also referred to as BsmBI) cleavage sites present in the original vector.
8. After the elution step, the linearized PCR product is ready for upcoming experimental steps.
Critical: To minimize the occurrence of gRNA-free colonies on negative control LB agar plates, utilize a maximum of 1 ng of lentiGuide-Puro per PCR reaction (see Troubleshooting 1). Additionally, to compensate for potential losses during purification, conduct a total of 10 PCR reactions.
Note: Scale up the number of reactions whenever there is a loss during consecutive purification steps.
9. Subsequently, for product size validation, 1) pool the eluted PCR-linearized lentiGuide-Puro into a single tube, and 2) mix the PCR linearization product with gel loading dye purple 6× to a final concentration of 1×. Then, run 5 μL of the validation reaction alongside uncut lentiGuide-Puro on a 0.8% (w/v) agarose gel cast in 1× TAE buffer with SafeView Classic dye (Figure 1, gel image on the left).
Figure 1. lentiGuide-Puro PCR linearization. The gel image on the left displays the result of the PCR linearization product on a 0.8% (w/v) agarose gel, with the last lane demonstrating 100 ng of uncut lentiGuide-Puro backbone as a control. As a DNA ladder, 1 kb plus DNA ladder was used. The gel image on the right shows the column-purified vector for pre-cloning after Esp3I (BsmBI) restriction digestion.
10. Following the pooling of PCR products and column purification of the lentiGuide-Puro backbone, which serves as the template for linearization, incubate with the Esp3I (BsmBI) restriction enzyme at 37 °C for 4 h.
Note 1: Initial optimization experiments employed a 1 h digestion, with 4 h ultimately identified as the optimal condition in our experimental setup.
Note 2: This procedure was intended to diminish the colony-forming potential of the intact lentiGuide-Puro vector used in the PCR-based linearization.
Critical: FastDigest Esp3I (BsmBI) enzyme requires dithiothreitol (DTT) and should be prepared freshly (see Troubleshooting 1). Reaction components for restriction digestion comprise 14 μL of nuclease-free water, 2 μL of 10× FastDigest Buffer, 1 μL of DTT (20 mM), 1 μL of FastDigest enzyme Esp3I (BsmBI), and 1 μg of the PCR product. Mix all the reaction components on ice, followed by incubation at 37 °C for 4 h.
11. Inactivate the restriction enzyme by incubating the reaction at 65 °C for 15 min.
12. Subsequently, repurify the linearized Esp3I (BsmBI) digested PCR product using NucleoSpin Gel and PCR clean-up Kit and verify for purity and integrity on 0.8% (w/v) agarose (as described in step A9) (Figure 1, gel image on the right). All modifications made at this stage should successfully eliminate factors affecting impurity.
B. PCR amplification and purification of the gRNA library
Note 1: The gRNA pool was improved upon the nuclear proteins gRNA sub-pool library (Addgene #51047 Nuclear Proteins gRNA sub-pool Library): an expanded list of genes targeting epigenetic regulators, transcription factors, and nuclear factors (labeled as EpiTransNuc) was assembled by adding validated gRNA sequences from various libraries, including Sabatini, Brunello, GecKO, and Toronto v3, all available from Addgene. For each gRNA, forward (F) and reverse (R) sequences were appended to both ends of the gRNA sequences. Oligonucleotide sequences consist of a 5' universal flanking sequence (TATCTTGTGGAAAGGACGAAACACCG) and a 3' universal flanking sequence (GTTTTAGAGCTAGAAATAGCAAGTTAAAAT). Oligonucleotides featuring a 5' and 3' extension compatible with the Esp3I (BsmBI) enzyme were synthesized in array format by CustomArray (Genscript).
Note 2: The EpiTransNuc library was designed so that each gene is targeted by 10 distinct gRNAs, covering a total of 4,072 genes. Additionally, 100 non-targeting control gRNAs were included in the library composition [retrieved from Addgene #51048 Control targets (most diverse) gRNA sub-pool library].
1. To transition from the gRNA oligonucleotide library to the PCR library, amplification was performed using NEBNext high-fidelity PCR master mix. The reaction was setup using the following primer pair: GTAACTTGAAAGTATTTCGATTTCTTGGCTTTATATATCTTGTGGAAAGGACGAAACACC (forward primer) and ACTTTTTCAAGTTGATAACGGACTAGCCTTATTTTAACTTGCTATTTCTAGCTCTAAAAC (reverse primer).
2. Set up a 6× gRNA oligonucleotide amplification reaction (Table 4) by following the thermocycling conditions shown in Table 5. Prepare the master mix in a 1.5 mL Eppendorf tube, then transfer it into 0.2 mL PCR tubes.
Table 4. Oligonucleotide library amplification
Reagents Final concentration Volume required (μL)
NebNext, HF, PCR MM, 2× 1× 12.5
Pooled oligonucleotide library template 0.04 ng/μL 1
Oligonucleotide forward primer 0.5 μM 1.25
Oligonucleotide reverse primer 0.5 μM 1.25
Ultra-pure water 9
Total 25
Table 5. Thermocycling conditions for the PCR reaction
Step Temp. (°C) Duration No. of cycles
Initial denaturation 98 30 s 1
Denaturation 98 10 s
20
Annealing 63 10 s
Extension 72 15 s
Final extension 72 2 min 1
Hold 4 ∞ -
3. After the amplification, 5 μL of the PCR product was electrophoresed on a 2.5% (w/v) agarose gel for validation (as described in step A9) (Figure 2).
Note: The PCR amplicon is observed at approximately 140 bp. Notably, the 120 bp primer dimer reaction, highlighted as a concern in the study by Joung et al. [11], was not detected in our samples. To mitigate potential skewing in gRNA representation, PCR amplification was restricted to 20 cycles (see Troubleshooting 5) in accordance with recommendations by Joung et al. [11].
Figure 2. PCR amplification of gRNA oligonucleotides. The oligonucleotides synthesized through a service provider were amplified using long primer pairs suggested by Joung et al. [11], and a small amount (5 μL) was analyzed on a 2.5% (w/v) agarose gel for validation purposes. A 1 kb plus DNA ladder was used. The first well represents the library PCR product, while the second well serves as the negative control (water as input).
4. gRNA PCR library was purified with the NucleoSpin Gel and PCR clean-up kit, following the protocol provided by the manufacturer.
Note: In line with the approach used for the linear vector preparation, the gel extraction step was omitted during the preparation of the gRNA PCR library to ensure purity.
C. Circular polymerization extension cloning (CPEC)
Note: Components of CPEC protocol comprise NEBNext HF PCR master mix, previously PCR-linearized lentiGuide-Puro backbone (A), PCR-amplified gRNA library (B), and distilled water.
1. Use 100 ng of lentiGuide-Puro backbone while utilizing only 16.9 ng of PCR amplified and purified gRNA library (Table 6). Limit the CPEC reaction to four cycles and perform at least 10 separate reactions following the thermocycling conditions shared in Table 7. Include negative controls involving the same setup without gRNA library addition.
Table 6. Circular polymerization and extension cloning reaction
Reagents Final concentration Volume required (μL)
NebNext, HF, PCR MM, 2× 1× 12.5
lentiGuide-Puro (Esp3I (BsmBI)) 1 M 100 ng (varies)
PCR product (gRNA library) 10 M 16.9 ng (varies)
ddH2O varies
Total 25
Table 7. Thermocycling conditions for the CPEC reaction
Step Temp. (°C) Duration No. of cycles
Initial denaturation 98 30 s 1
Denaturation 98 10 s
4
Primer annealing 72 5 min
Extension 72 5 min
Final extension 72 5 min 1
Hold 4 ∞ -
2. Following this, purify the CPEC products using isopropanol, as recommended by Joung et al. [11]. In this procedure, mix the CPEC product with isopropanol at a 1:1 ratio. Subsequently, add NaCl solution to achieve a final concentration of 50 mM.
Note: This precipitation step is intended to concentrate the library prior to bacterial transformation and to reduce the salt concentration from prior reactions, which could adversely impact electroporation efficiency (see Troubleshooting 4).
3. Vortex the reaction and incubate at room temperature for 15 min. Centrifuge the CPEC product at 15,000× g for 15 min at room temperature.
Critical: At this stage, the pellet may be difficult to visualize. Carefully aspirate the supernatant from the side opposite to where the pellet is located to avoid accidentally aspirating the CPEC product (see Troubleshooting 2). Use the vacuum aspiration system at the lowest setting.
4. Add ice-cold 80% ethanol (150 μL) without disturbing the pellet and centrifuge samples at 11,000× g for 5 min at 4 °C. Aspirate the supernatant carefully and repeat this step one more time. Remove any residual ethanol and air-dry samples for 1 min.
5. Incubate the reaction at 55 °C for 10 min with a TE buffer.
6. Determine the concentration using a NanoDrop2000 spectrophotometer.
D. Electroporation of the CPEC product
1. Transfer the CPEC reaction (600 ng of total gRNA plasmid library) to Endura electrocompetent bacteria. At this step, carry out three separate electroporations, each one with 25 μL of electrocompetent bacteria.
2. Perform electroporation by following the EC1 program on Bio-Rad MicroPulser.
Note 1: In this setting, the convenient cuvette size is 0.1 cm, the voltage is 1.8 kV, and the number of pulses is 1 (6 ms). Electroporation conditions are based on 10 microfarads and 600 ohm resistance.
Note 2: Use 2 mL of recovery medium.
3. Following this, pool bacteria and incubate for 75 min at 225 rpm and 37 °C conditions.
4. Spread the bacteria carrying the target library onto two prewarmed LB agar bioassay dishes (500 cm2) and incubate for 18 h at 37 °C culture conditions (Figure 3).
Note 1: Simultaneously, in order to determine the library coverage, spread the bacteria in different dilutions (1/10,000, 1/1,000, and 1/10) onto ampicillin-containing (1:1,000) agar plates.
Note 2: In Figure 3, there were 1,761 CPEC (+) colonies in 1/1,000 dilution, whereas there were only 22 colonies in CPEC (-) plate (see Troubleshooting 3).
Note 3: Our library coverage was calculated as ~43× using the following formula:
Coverage = (CPEC (+) colony number - CPEC (-) colony number) * Dilution factor/Library size
According to our results, the formula applies as (1761 - 22 = 1739) * 1,000 (dilution factor) = 1,739,000. Divide the result by the library size; in our case, this is 40,820 gRNAs.
Figure 3. Electroporation application of commercial Endura bacteria. Circular polymerase extension cloning (CPEC) (+) and CPEC (-) products in varying dilutions on 10 cm Petri dishes (left). CPEC (+) product spread onto two LB agar bioassay dishes (representative image on the right).
5. After 18 h, gently scrape the electrocompetent bacteria from LB agar bioassay dishes by using a bio-spreader with 20 mL of cold LB liquid medium per dish. Centrifuge (previously set to 4 °C) Falcon tubes with bacteria for 30 min at 3,214× g and, after removal of supernatants, weigh pellets.
6. Purify the products with an endotoxin-free HiSpeed Plasmid DNA Maxi kit.
Data analysis
The gRNA distribution in the library was confirmed by Next-generation sequencing (NGS) using HiSeq X with 2× 150 bp paired-end reads, providing a sequencing depth of 6 Gb per sample. NGS library was constructed following the protocols and reagents described by Joung et al. [11]. The raw sequencing data was retrieved and processed using the count_spacers.py script in Python v2.7.17 [11].
Validation of protocol
We validated the process using the count_spacers.py code along with additional analyses. To run count_spacers.py, three essential inputs are required: 1) the .fastq file containing sequencing data (Figure 4), 2) a .csv file with gRNA sequence information, and 3) a key sequence (CGAAACACC) that facilitates the identification of gRNAs within the sequencing reads. The key sequence may be tailored based on the specific sequence composition of the library.
Figure 4. Subset of the NGS sequencing .fastq file used in the analysis
When necessary, the .fastq files were trimmed to remove adapter sequences using the trimming algorithm like Cutadapt. The trimmed .fastq files were analyzed using the count_spacers.py code (https://github.com/fengzhanglab/Screening_Protocols_manuscript/blob/master/count_spacers.py), which generated two output files: 1) a text file containing analysis statistics (Figure 5) and 2) a .csv file listing the read counts for each gRNA (Table 8).
Figure 5. Statistics.txt output file from count_spacers.py code
Table 8. Read counts for the top 10 gRNAs from .csv file
gRNA Sequence Read count
TCCGCGCCTTCGCCTACACC 1,818
TGCTGTCCACCGCTCCTCCC 1,553
TGCCTGTCCTGTGTCAAGTC 1,533
TGAACTCGTCCAGCACCGCC 1,505
AGCCGCGCCTCACCGGGTGC 1,447
TCACTCACCTGCATCTGCCC 1,447
TCTGCCTTGTTCCCTGCCTG 1,446
TGTCTGCTGCTCCTGCCTTT 1,416
GTCCCAGTTCTCCGCCCTCC 1,407
TTCCTCCTCGCTCTCCTCTC 1,402
To visualize and assess the distribution of gRNAs, a density plot was generated using the log2-transformed counts of the gRNAs (Figure 6). The density plot was created using R (v4.2.2) and the ggplot2 package (v3.5.1). The analysis revealed that over 90% of the gRNAs were represented within a 10-fold range of read counts, indicating a highly uniform representation across the library. This suggests minimal over- or under-representation of individual gRNAs, ensuring the integrity of the library for downstream applications.
Figure 6. Density plot depicting the gRNA representation in EpiTransNuc library. The distribution of gRNAs in the library was verified using NGS. The bars in the graph illustrate the log2-transformed read counts for each gRNA.
General notes and troubleshooting
General notes
1. To prevent inter-colony competition that may result in skewing of gRNA library distribution, a solid culture amplification rather than a liquid one is recommended [11].
2. Multi-fragment assembly can be accomplished via CPEC methodology so that at least two gRNAs can be cloned tandemly in one vector backbone in order to target different genes.
Troubleshooting
Problem 1: High number of colonies on the negative control agar plates.
Possible cause: Recovery of backbone, incomplete restriction of uncut plasmid.
Solution 1: In the PCR-based linearization, usage of 1 ng of backbone per reaction is strictly recommended.
Solution 2: For 1 μg of PCR-linearized product, usage of 1 μL of Esp3I (BsmBI) is quite significant; do not reduce the enzyme amount.
Solution 3: FastDigest Esp3I (BsmBI) enzyme requires DTT in the reaction. DTT should be prepared and added freshly.
Problem 2: Aspiration of CPEC products after isopropanol precipitation step.
Possible cause: Pellet is loose or difficult to see.
Solution 1: Perform a second round of centrifugation.
Solution 2: Use GlycoBlue, a reagent that comprises a blue dye covalently conjugated to glycogen. It serves as a nucleic acid co-precipitant, with the dye facilitating enhanced visualization of the nucleic acid pellet.
Problem 3: Insufficient number of colonies on the positive agar plates, low transformation efficiency.
Possible cause: Gel extraction step interferes with the CPEC reaction and mitigates positive clones. Moreover, check your electrocompetent cells with an intact, commercial plasmid.
Solution: Skip gel extraction steps wherever it is feasible and scale up the number of electroporation reactions.
Problem 4: Problems or arcing during electroporation.
Possible cause: Salts coming from buffers, higher ionic strength of samples.
Solution: Perform isopropanol precipitation, desalt as much as possible before electroporation, and ensure that there are no air bubbles in the electroporation cuvette.
Problem 5: Insufficient representation of the gRNA library.
Possible cause: PCR-based bias
Solution: Minimize the number of PCR cycles during oligonucleotide amplification to prevent potential biases; limit PCR cycles to 20.
Acknowledgments
The authors acknowledge Izmir Biomedicine and Genome Center (IBG) for providing financial and administrative support. This research was funded by The Scientific and Technological Research Council of Türkiye (TÜBİTAK) with grant number 119Z540. Bengisu Dayanc was supported by YÖK 100/2000 PhD Scholarship and TÜBİTAK-BİDEB 2211/C National PhD Scholarship programs. Sude Eris was supported by TÜBİTAK-BİDEB 2210/A National MSc/MA scholarship program. Serif Senturk acknowledges support from the Turkish Academy of Sciences (TUBA GEBIP 2017) and the Science Academy (BAGEP 2019). We acknowledge Dr. Minoo Karimi (University of Montreal) and Dr. Ece Cakiroglu (Dokuz Eylul University) for technical help throughout the construction of the gRNA library.
Competing interests
The authors declare no competing interests.
References
Shalem, O., Sanjana, N. E. and Zhang, F. (2015). High-throughput functional genomics using CRISPR–Cas9. Nat Rev Genet. 16(5): 299–311.
Zhang, Z., Wang, H., Yan, Q., Cui, J., Chen, Y., Ruan, S., Yang, J., Wu, Z., Han, M., Huang, S., et al. (2023). Genome-wide CRISPR/Cas9 screening for drug resistance in tumors. Front Pharmacol. 14: e1284610.
Chan, Y. T., Lu, Y., Wu, J., Zhang, C., Tan, H. Y., Bian, Z. x., Wang, N. and Feng, Y. (2022). CRISPR-Cas9 library screening approach for anti-cancer drug discovery: overview and perspectives. Theranostics. 12(7): 3329–3344.
Sanson, K. R., Hanna, R. E., Hegde, M., Donovan, K. F., Strand, C., Sullender, M. E., Vaimberg, E. W., Goodale, A., Root, D. E., Piccioni, F., et al. (2018). Optimized libraries for CRISPR-Cas9 genetic screens with multiple modalities. Nat Commun. 9(1): 5416.
Quan, J. and Tian, J. (2009). Circular Polymerase Extension Cloning of Complex Gene Libraries and Pathways. PLoS One. 4(7): e6441.
Aslanidis, C. and de Jong, P. J. (1990). Ligation-independent cloning of PCR products (LIC-PCR). Nucleic Acids Res. 18(20): 6069–6074.
Li, M. Z. and Elledge, S. J. (2007). Harnessing homologous recombination in vitro to generate recombinant DNA via SLIC. Nat Methods. 4(3): 251–256.
Gibson, D. G., Young, L., Chuang, R. Y., Venter, J. C., Hutchison, C. A. and Smith, H. O. (2009). Enzymatic assembly of DNA molecules up to several hundred kilobases. Nat Methods. 6(5): 343–345.
Wegner, M., Husnjak, K. and Kaulich, M. (2020). Unbiased and Tailored CRISPR/Cas gRNA Libraries by Synthesizing Covalently-closed-circular (3Cs) DNA. Bio Protoc. 10(1): e3472.
Quan J, Tian J. (2014). Circular Polymerase Extension Cloning. In: Valla S, Lale R (Eds.). DNA Cloning and Assembly Methods. Humana Press, Totowa, NJ, pp 103–117.
Joung, J., Konermann, S., Gootenberg, J. S., Abudayyeh, O. O., Platt, R. J., Brigham, M. D., Sanjana, N. E. and Zhang, F. (2017). Genome-scale CRISPR-Cas9 knockout and transcriptional activation screening. Nat Protoc. 12(4): 828–863.
Article Information
Publication history
Received: Sep 25, 2024
Accepted: Dec 8, 2024
Available online: Dec 31, 2024
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Cancer Biology > General technique
Molecular Biology > DNA > DNA cloning
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5,184 | https://bio-protocol.org/en/bpdetail?id=5184&type=0 | # Bio-Protocol Content
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This is an In Press version of the protocol that has not yet been assigned to an issue.
Peer-reviewed
Quantifying Bacterial Chemotaxis in Controlled and Stationary Chemical Gradients with a Microfluidic Device
Adam Gargasson
CD Carine Douarche
Peter Mergaert
HA Harold Auradou
In Press, Available online: Dec 31, 2024
DOI: 10.21769/BioProtoc.5184 Views: 82
Reviewed by: Hsih-Yin Tan Anonymous reviewer(s)
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Abstract
Chemotaxis refers to the ability of organisms to detect chemical gradients and bias their motion accordingly. Quantifying this bias is critical for many applications and requires a device that can generate and maintain a constant concentration field over a long period allowing for the observation of bacterial responses. In 2010, a method was introduced that combines microfluidics and hydrogel to facilitate the diffusion of chemical species and to set a linear gradient in a bacterial suspension in the absence of liquid flow. The device consists of three closely parallel channels, with the two outermost channels containing chemical species at varying concentrations, forming a uniform, stationary, and controlled gradient between them. Bacteria positioned in the central channel respond to this gradient by accumulating toward the high chemoattractant concentrations. Video-imaging of bacteria in fluorescent microscopy followed by trajectory analysis provide access to the key diffusive and chemotactic parameters of motility for the studied bacterial species. This technique offers a significant advantage over other microfluidic techniques as it enables observations in a stationary gradient. Here, we outline a modified and improved protocol that allows for the renewal of the bacterial population, modification of the chemical environment, and the performance of new measurements using the same chip. To demonstrate its efficacy, the protocol was used to measure the response of a strain of Escherichia coli to gradients of α-methyl-aspartate across the entire response range of the bacteria and for different gradients.
Key features
• The protocol is based on a previously proposed system [1] that we improved for higher throughput.
• Setup allowing a rapid quantification of motility and chemotaxis responses.
• Seventeen hours were required from the start of an E. coli culture to the measurements to obtain the chemotactic velocity under various chemical conditions.
Keywords: Microfluidics Chemotaxis Bacteria Flow free Microscopy Image analysis
Graphical overview
Schematic illustration of the three-channels chip architecture and its use with bacteria
Background
Bacterial chemotaxis is an important area of research that spans multiple fields. Examples include (i) microbiology, in which it helps to explain fundamental aspects of bacterial behavior, signaling pathways, and cellular processes; (ii) infectious diseases, where insights into how pathogenic bacteria use chemotaxis to locate and infect host tissues can inform the development of new treatments and preventive measures; (iii) environmental microbiology, in which chemotaxis is crucial for bacterial movement in natural environments, influencing nutrient cycling, bioremediation, and the functioning of microbial ecosystems; (iv) ecology and evolution, in which chemotaxis affects microbial interactions, competition, and cooperation within communities, providing insights into microbial ecology and the evolutionary dynamics of these behaviors; and (v) agriculture, where understanding how soil bacteria use chemotaxis to interact with plant roots can improve agricultural practices, enhance crop growth, and lead to the development of biofertilizers or biopesticides.
It is accepted that the chemotactic drift takes place at a velocity
v
c
→
dependent on the detected concentration c and local gradient
∇
c
→
of the chemoattractant according to the following equation [2,3]:
v
c
→
=
χ
(
c
)
∇
c
→
(1)
where
χ
(
c
)
is the chemotactic susceptibility that depends on the local concentration in the bacterial vicinity. Numerous assays have been used to determine the chemotactic response of bacteria. The capillary tube assay is a classical method developed in the late XIXe century [4], which involves using a capillary tube filled with a chemoattractant solution. The tube is placed in a bacterial suspension, and the number of bacteria that enter the tube is counted. Some assays use the displacement of bacteria on soft agar plates containing the chemoattractant to measure chemotactic velocity. Several microfluidic solutions have been developed [5], including a recently introduced high-throughput method [6]. These assays can provide quantitative insights into the chemotactic behavior of bacteria. However, in order to determine vc, it is necessary to control the stationarity of the gradient and maintain shear-free conditions.
To this end, our microfluidic device uses diffusion through agar to generate a stable and controlled concentration and gradient and allows measuring precisely the vc for this environment. In practice, the gradient is established between parallel channels separated by a distance L, each containing a concentration cmin and cmax. Its value is
∇
c
=
c
m
a
x
-
c
m
i
n
L
and the average concentration of chemoattractant in which the bacteria swim is
c
=
c
m
a
x
+
c
m
i
n
2
. Bacteria are introduced into the device, and their movement in response to the gradient is observed using microscopy. Bacteria swim toward the chemoattractant and gather on the channel wall closest to the source of the chemoattractant. Once the stationary regime is reached, the concentration of bacteria in this region decays exponentially, with a decay length:
λ
=
D
v
c
(2)
This results from the competition between the advection of bacteria toward the source of chemoattractant at velocity vc and their active diffusion, which is characterized by a diffusion coefficient D. Here, the diffusion coefficient D is obtained by analyzing bacterial trajectories. This method enables the quantitative determination of chemotactic velocity and its dependence on the average concentration and its gradient. The chemotactic coefficient can be determined from the relation (see Eq. 1):
χ
(
c
)
=
L
v
c
c
m
a
x
-
c
m
i
n
(3)
In its initial version, published by Ahmed et al. [1], the test required one microfluidic device per trial, making the method very time-consuming. Our protocol allows multiple trials to be run on the same chip, which may allow the method to be automated and enable continuous measurements. The method requires specialized equipment and expertise, which are presented here in detail. We illustrate the protocol using an Escherichia coli strain placed in α-methyl-aspartate gradients.
Materials and reagents
Biological materials
1. E. coli RP437 [7] (CGSC: #12122 transformed with the plasmid pZA3R-YFP carrying a chloramphenicol resistance marker and a yfp gene) stored in 25% glycerol in a -80 °C freezer for long-term conservation
Reagents
1. M9 salts (MP Biomedicals, catalog number: SKU 113037012-CF), dissolved in water at 22% (11 g in 50 mL)
2. Casamino acids (VWR, catalog number: ICNA113060012), dissolved in water at 5% (2.5 g in 50 mL)
3. D-(+)-Glucose (Sigma-Aldrich, catalog number: G7021-1KG), dissolved in water at 20% (10 g in 50 mL)
4. MgSO4·7H2O (Sigma-Aldrich, catalog number: 63138-250G), dissolved in water at 1 M (1.2 g in 50 mL)
5. KH2PO4 (Sigma-Aldrich, catalog number: P5655-500G), dissolved in water at 1 M (3.4 g in 25 mL water)
6. K2HPO4 (Sigma-Aldrich, catalog number: P3786-500G), dissolved in water at 1 M (4.3 g in 25 mL water)
7. CaCl2·2H2O (Sigma-Aldrich, catalog number: C3306-100G), dissolved in water at 1 M (36.8 mg in 50 mL)
8. Sodium lactate (Sigma-Aldrich, catalog number: L7022-10G), dissolved in water at 100 mM (1.1 g in 10 mL)
9. EDTA (Sigma-Aldrich, catalog number: EDS-100G), dissolved in water at 10 mM (292 mg in 10 mL)
10. L-methionine (Sigma-Aldrich, catalog number: M5308-25G), dissolved in water at 100 μM (149 mg in 1 L)
11. Chloramphenicol (Sigma-Aldrich, catalog number: C0378-5G)
12. Bacto agar (BD-Difco, catalog number: 214010)
13. PDMS components (Ellsworth, Dow Sylgard 184, GMID: 1673921)
a. Monomer
b. Curing agent
14. Ethanol 95% (Fisher Chemical, catalog number: E/0500DF/P21)
15. α-methyl-aspartate (MedChem, catalog number: HY-W142119)
Solutions
1. 100 mM KHPO4 buffer, pH 7 (see Recipes)
2. M9G medium (see Recipes)
3. Motility buffer (see Recipes)
4. PDMS (see Recipes)
5. 3% agar gel (see Recipes)
6. 70% ethanol (see Recipes)
7. Chloramphenicol stock solution (see Recipes)
Recipes
1. 100 mM KHPO4 buffer, pH 7
Reagent Final concentration Amount
KH2PO4 (1 M) 38.5 mM 1.54 mL
K2HPO4 (1 M) 61.5 mM 2.46 mL
H2O n/a 36 mL
Total n/a 40 mL
2. M9G medium
Reagent Final concentration Amount
M9 salts (220 g/L) 11.1 g/L 25 mL
D-(+)-Glucose (200 g/L) 4 g/L 10 mL
Casamino acids (50 g/L) 1 g/L 10 mL
MgSO4·7H2O (100 mM) 2 mM 10 mL
CaCl2·2H2O (5 mM) 100 μM 10 mL
H2O n/a 435 mL
Total n/a 500 mL
3. Motility buffer
Reagent Final concentration Amount
KHPO4 buffer (100 mM, pH 7.0) 10 mM 1 mL
Sodium lactate (1 M) 10 mM 100 μL
EDTA (100 mM, pH 10.0) 100 μM 10 μL
L-Methionine (1 mM) 1 μM 10 μL
H2O n/a 8.88 mL
Total n/a 10 mL
4. PDMS
Reagent Final concentration Amount
Monomer 90% 90 g
Curing agent 9% 9 g
Total n/a 99 g
5. 3% agar gel
Reagent Final concentration Amount
Bacto Agar 30 g/L 300 mg
H2O MilliQ n/a 10 mL
Total n/a 10 mL
6. 70% ethanol
Reagent Final concentration Amount
Ethanol (95%) 70% 350 mL
H2O n/a 125 mL
Total n/a 475 mL
7. Chloramphenicol stock solution
Reagent Final concentration Amount
Chloramphenicol 25 g/L 250 mg
70% ethanol n/a 10 mL
Total n/a 10 mL
Store at -20 °C.
Laboratory supplies
1. Round-bottom double position 14 mL tubes (Falcon, catalog number: 352057)
2. 5 mL pipettes (Costar, Stripette, catalog number: 4487)
3. Micropipette tips (Gilson, D1000, catalog number: F167104)
4. Glass Petri dish D 120 mm, H 20 mm (Rogo Sampaic, catalog number: BRB005)
5. Scalpels (Swann Morton, catalog number: 0501)
6. Tubing (Darwin, Tygon, catalog number: LVF-KTU-13)
7. Safe-lock microtubes 2.0 mL (Eppendorf, catalog number: 0030120094)
8. Microvalve (Cluzeau CIL, catalog number: P-782)
9. 2 mm diameter sterile disposable biopsy punches (INTEGRA Miltex, catalog number: 33-31)
10. 2.5 mL syringes (Hamilton, Gastight 1002, catalog number: 81420)
11. Microscope glass slides 76 × 26 × 1 mm (Brand, catalog number: 474743)
12. Microscope glass slides 75.5 × 51.5 × 1.0 mm (Knittel, catalog number: VY11300051075.01)
13. Disposable plastic beaker 400 mL (Azlon, catalog number: BBBPB0400P) or paper cup
Equipment
1. Microscope (Leica, model: DMI 6000B, catalog number: 11888941)
a. Base (Leica, model: CTR6000, catalog number: 11888821)
b. Lens (Leica, model: HC PL FLUOTAR L 20×, catalog number: 11506243)
c. Fluorescence cube (Leica, model: Fluorescence Filter, Blue, I3, catalog number: 11513878): excitation filter: 450–490 μm; dichromatic mirror: 510 μm; suppression filter: 515 μm
d. Fluorescent light source (Leica, model: EL 6000, catalog number: 11504115)
e. Controller (Leica, model: IV/2013, catalog number: 11505180)
2. Digital CMOS camera (Hamamatsu, model: Orca-Flash4.0 V3, product number: C13440-20CU) mounted on the microscope
3. Syringe pump (Cetoni)
a. Base module (Cetoni Base 120, model: NEM-B100-01 F)
b. Three dosing units (Cetoni, Dosingmodule 14:1, model: NEM-B101-02 D)
Less sophisticated pumps can be used, including self-made pumps, see this link for example
4. Spectrophotometer (Eppendorf, model: D30, catalog number: 6133000001)
5. Pipette controller (Integra, model: PIPETBOY pro, catalog number: 156403/156401)
6. Biological safety cabinet (Telstar, model: Bio II Advance 3, catalog number: 523913)
7. Incubator shaker (Eppendorf, model: New Brunswick Innova 40R, catalog number: M1299-0086)
8. Autoclave (Advantage-Lab, model: AL02-01-100)
9. Oven [Labnet, catalog number: I5110(A)-230V]
10. Void Pump (KNF Lab, Laboport N816.3 KN.18, catalog number: 03533752)
11. Mini centrifuge (Sigma, model: 1-14, catalog number: 10014)
12. Stirrer hotplate (Fisher Scientific, catalog number: FB15001)
13. Semi-micro balance [Sartorius, model: CP(A)225D]
14. Precision balance (Sartorius, model: CP420S)
15. Vacuum desiccator (Bel-art, catalog number: F42022-0000)
16. Vortex mini mixer (Crystal LabPro, model: VM-03RUW)
17. -80 °C ultra-low temperature freezer (New Brunswick, New Brunswick Innova U101, model: U101-86)
18. Refrigerator: -20 °C freezer on top (Proline, DD133, catalog number: 7605749)
Software and datasets
1. NemeSys (v. 2016.06.14)
2. Leica Application SuiteX (v. 3.4.7)
3. Fiji (v. 1.52; https://imagej.net/software/fiji/downloads)
4. Python (v. 3.9; https://www.python.org/)
5. Inkscape (v. 1.3.2; https://inkscape.org/)
Procedure
A. Obtaining the suspensions of E. coli used to validate the protocol
1. Add 5 mL of M9G medium and 5 μL of chloramphenicol stock solution into a double-position tube and vortex.
2. Inoculate the M9G medium with a small piece of ice scraped from the -80 °C glycerol stock of the E. coli strain and incubate overnight at 30 °C and 240 rpm.
3. When OD600 ~0.1, transfer 1 mL of the culture into a microtube and centrifuge it at 10,000 rpm (7,378× g) for 5 min. Remove the supernatant and resuspend the bacterial pellets in 1 mL of Milli Q water. Centrifuge the microtube again at 10,000 rpm (7,378× g) for 5 min. Remove the Milli Q water and resuspend the bacteria in 1 mL of motility buffer.
4. Measure the OD600 with the spectrophotometer and adjust the suspension to an OD600 ~0.08 by adding the appropriate volume of motility buffer. The scarcity of carbon source maintains the OD600 for more than 3 h, and the suspension is still motile after 1 day in a closed microtube.
B. Fabrication of the microfluidic chip
1. Negative imprint of channels in photosensitive resin on a wafer for PDMS molding.
The manufacturing of the wafer requires a photolithography platform that can be found in nanotechnology laboratories. We used the C2N (Centre des Nanosciences et des Nanotechnologies) facilities to realize the imprint. The height of the channels is set by the thickness of the SU8 resin spin-coated onto a 4-inches silicon wafer. The experiments reported here were carried out with 117 μm high channels. This method can achieve smaller heights by adjusting the spin-coater rotation speed and resin viscosity. When the resin is exposed to UV light, it polymerizes, attaches to the surface of the wafer, and becomes solid. To fix the resin where the channels will be present, a photomask must be placed between the UV lamp and the wafer on which the resin is deposited. The photomask must be printed using a very high-resolution printer to achieve good spatial resolution. Selba produced our photomask with a 25,400 dpi resolution. The channels printed on the photomask were designed on Inkscape, and the SVG file was provided to the company. The SVG file can be downloaded here. Several geometries are available in the file and can be used to make the microfluidic cell. We have used the one where the channels are 600 μm wide and 200 μm apart.
2. Molding of the PDMS channels.
a. Mix 100 mL of PDMS monomer and curing agent with a spatula in a 100 mL beaker for 1–2 min. Remove the bubbles generated during mixing by placing the beaker in a vacuum chamber.
b. Cover the bottom of a Petri dish with a plastic film and aluminum foil. This will make it easier to release the molding. Carefully place the wafer inside and pour the degassed PDMS onto it. If there are any remaining bubbles, use a scalpel to remove them. Place the Petri dish in an incubator for at least 3 h at 60 °C allowing the polymerization of the PDMS.
c. Unmold the PDMS and cut out the area containing the channels. The side on which the channels are molded can be protected from dust by placing the piece of PDMS on a cleaned glass surface. Eventual dust deposited on the PDMS surface should be removed by pressing and removing a piece of tape on the surface of the PDMS.
d. To connect the chip to the pump and the reservoirs, make 2 mm radius holes with a biopsy punch at the enlarged extremities of the channels. Then, flexible tubings are inserted into the holes. The three output tubings, at one side of the channels, are connected to the pump; the three inlet tubings at the other side are dipped in a 100 mL water-filled Erlenmeyer.
The whole procedure is shown in Video 1.
Video 1. Preparing the PDMS molding
3. Preparation of the agar layer.
a. Place a 15 mL Falcon tube containing 10 mL of 3% agar gel in a boiling water bath until fully dissolved.
b. To obtain a thin flat surface of an agar gel that will be subsequently used to contact the PDMS layer, use the following procedure: (i) Make a flat surface out of PDMS: pour PDMS into a Petri dish, then degas and place in the incubator as above before cutting a 6 × 6 cm piece. (ii) Place two 75 × 26 mm microscope slides on the PDMS surface. Make sure they are separated by 5 cm, with their long sides as parallel as possible. They act as spacers, and their thickness determines the final thickness of the agar layer. Use glass slides 1 mm thick; this thickness can be increased by layering multiple microscope slides. (iii) Pour a few milliliters of molten agar on the surface, then cover it with a 75 × 50 mm microscope slide. This operation is done by sliding the microscope slide slowly over the spacers to allow the drops to spread and evenly fill the space between both spacers. Then, place the ensemble in the fridge for 10 min.
c. This operation produces a gel surface of agar, which needs to be transferred onto a microscope slide. First, take off the microscope slide placed on top of the agar. Then, scrutinize the agar layer to ensure the absence of imperfections. Slide the agar from the PDMS surface onto a new 75 × 50 mm microscope slide. Video 2 shows the whole procedure.
Video 2. Preparing the agar layer
4. Making the microfluidic chip with the PDMS mold and the agar surface (Figure 1).
a. First, fill the tubings with water. To do so, use the pump or create a gravity flow with the Erlenmeyer. It is important to avoid leaving bubbles in the tubes. They may prevent the proper fluid flow in the channels of the microfluidic chip.
b. Wet the agar layer with a few drops of water, then place the PDMS molding with the channels facing down on top of the agar layer. Press gently to avoid bubbles still trapped between the PDMS and the agar. The assembly can then be positioned on the microscope's moving stage.
c. Withdraw water with the pump set at a 10 μL/min flow rate to remove the water layer between the PDMS molding and the agar. The pump can be used with the NemeSys software provided with the device. If the water layer is not removed within 1 min, use a syringe to suck the water manually by placing the syringe on the side of the chip. Once the water layer is removed, the PDMS and agar parts are in contact, and water flows equally through all three channels. Visualize the flow in brightfield mode by focusing the microscope on the channels at 10× magnification and taking advantage of particle impurities present in the liquids. There should be no particle movement at the contact areas between the PDMS and agar.
d. The chip is ready for the experiment. Replace the Erlenmeyer filled with water with microtubes containing the chemoattractant and the bacteria. This is done by quickly removing the tubings from the Erlenmeyer and dipping them into the microtubes. Set the flow rate to 2 μL/min to purge the water from the tubings and fill them with the liquids.
e. When the liquids arrive in the microfluidic chip (this takes 10 min for 50 cm long tubings), reduce the flow to 1 μL/min. Wait for the concentration gradient to form between the two outer channels. The gradient is established as the chemoattractant diffuses through the agar. We advise waiting for 30 min until the gradient is stationary.
f. Adjust the microscope to view the central channel of the chip and focus on the fluid in the channel. Stop the flow in the central channel when the number of visible bacteria is constant. This takes less than 5 min.
Figure 1. Schematic diagram of the microfluidic device. Microtubes C1 and C2 contain a chemoattractant solution at two different concentrations C1 and C2. The central microtube contains the bacterial suspension. Microtubes C3 and C4 contain another pair of concentrations that can be studied after measurement with the first pair of microtubes. The top, middle, and bottom syringes contain the solutions withdrawn from the microtubes containing the chemoattractant at concentration C1 and the chemoattractant at concentration C2. The central syringe chemoattractant concentration is
c
1
+
c
2
2
due to diffusion across the agar gel and the bacterial suspension.
C. Setting the microscope to observe bacteria
1. Set the acquisition software to acquire with a 2 × 2 binning (this increases the image's brightness and eases the image treatment). Use a 20× magnification objective so that the field of view encompasses the entire width of the central channel. For a YFP-expressing strain, such as RP437, use an I3 fluorescence filter cube. Make the focus at the bottom of the channel. We found that 10 frames per second (fps) for 20 s is sufficient to perform the statistical analysis of the trajectories.
2. Start acquisition 20 min after step B4f.
3. Save the films as a LIF project (Leica Image File).
4. New gradients can be generated by changing microtubes A and C with microtubes containing new chemical concentrations. Repeat the protocol from step B4d. It is recommended to start with low concentrations and to increase them from one experiment to another to avoid bias due to the residual presence of chemoattractant.
Data analysis
The goal of the data analysis is to extract the accumulation distance value λ and the diffusion coefficient of the bacteria D from the tracks. Based on these two quantities, the chemotactic velocity will be calculated using Eq. (2). Section D explains how to extract the trajectories of the bacteria from the movies, while Section E provides insight into the method used to determine the diffusion coefficient and the accumulation distance.
A. Obtaining the positions and trajectories of bacteria
1. With Fiji, open the LIF project with the Bio-Format Importer and select the movies.
2. Tracking is then carried out using the TrackMate® plug-in [8]. Tracking determines the positions of bacteria and filters out motile and non-motile bacteria. We have identified three crucial stages in this operation:
a. It is necessary to give a dimension to the objects to be detected (Figure 2A). This parameter is set at 4 µm, slightly above the size of bacteria. TrackMate attributes a quality factor to each detected feature, which reflects its sharpness in the focal plane. Features with a low-quality factor are out of focus. To eliminate false detections, a threshold for the quality factor is selected. The distribution of quality factors is bimodal, and the threshold is defined between the two distributions (Figure 2A).
b. The second important stage is the reconstruction of the trajectories, which involves identifying the particles between two successive images. Set the tracker to 4 µm (the distance around which the algorithm searches for the bacteria in the next image) to guarantee a good trajectory reconstruction (Figure 2B).
c. Before saving the trajectories to a file, a filter is applied to the length of the trajectories. Only trajectories of more than 10 positions are saved (Figure 2C). This corresponds to trajectories longer than 1 s.
d. Save the track table as a CSV file.
Figure 2. Screenshots taken during film analysis to determine bacterial trajectories using the TrackMate plugin. (A) Window in which you can enter the size and quality factor for detecting bacteria on each of the images. (B) Window that opens when you need to enter the parameters that will be used to link the particles detected between two successive images and thus reconstruct its trajectory. (C) Window at which filters can be applied to the trajectories. We have chosen to keep only trajectories containing more than 10 positions.
B. Suppression of the non-motile bacteria from the tracks, determination of the accumulation length and diffusion coefficient from the trajectories, and determination of the chemotactic velocity
1. Treat the CSV files with a tabulator. We recommend using the Pandas library in Python. It is also possible to use Microsoft Excel. A simplified version of the Python code we used is available in File S1.
2. The CSV file lists the trajectories by index and gives the coordinates of the bacteria determined at time t for each trajectory xi(t),yi(t). From the position, determine the following: (i) the velocity components vix(t) (along the channel direction) and viy(t) (in the direction normal to the channel) of the bacteria of index i at time t by applying the formula:
v
i
x
(
t
)
x
i
(
t
+
δ
t
)
-
x
i
(
t
)
δ
t
and
v
i
y
(
t
)
=
y
i
(
t
+
δ
t
)
-
y
i
(
t
)
δ
t
, where δt is the time taken to capture two successive images; (ii) the average velocity
v
i
=
<
∥
v
i
→
(
t
)
∥
t
>
of each trajectory i, where:
∥
v
i
→
(
t
)
∥
=
v
i
x
(
t
)
2
+
v
i
y
(
t
)
2
, (iii) the average velocity
v
y
component along the y direction by averaging the velocity component viy(t) over time t and trajectory index i.
3. Non-motile bacteria are in the liquid, while others stick to the surface and are recorded during the image acquisition. As this sub-population shows no chemotaxis, it is necessary to remove these bacteria from the list of trajectories obtained previously before conducting a statistical analysis of the trajectories. This is done by removing the trajectories with an average velocity vi higher than a threshold of 5 μm/s.
4. Determination of the diffusion coefficient from the average value of and the persistence time τ of the bacteria trajectories.
a. The persistence time measures the time over which the orientation of a bacterium's trajectory remains identical [9]. Over longer times, the trajectory is equivalent to that of a random walker. This time is obtained from the velocity correlation function calculated as [3]:
c
y
(
∆
t
)
=
<
v
i
y
(
t
)
·
v
i
y
(
t
+
∆
t
)
>
i
,
t
(4)
For particles randomly exploring their environment, the correlation function decays like
C
y
(
∆
t
)
=
v
y
2
e
-
∆
t
τ
(5), where τ is the correlation time we look for and vy is the average velocity of all the particles along the y axis estimated in step B2 (Data analysis).
b. When the correlation function is divided by , the only remaining fitting parameter of Eq. (5) is τ, obtained by an exponential fit of the data in the range 0.5 s < Δt < 3 s.
c. The diffusion coefficient is
D
=
v
y
2
τ
[9].
5. Determination of the accumulation characteristic length λ from the positions of the bacteria in the channel.
a. Segment the image along the y-axis into 20 μm wide bins and measure the bacterial density by counting the number of spots inside each bin to finally get the density distribution.
b. The concentration of bacteria in the stationary regime decays exponentially and is described by the function
b
(
y
)
=
b
0
e
-
y
-
y
0
λ
, where y0 is the upper boundary location, b0 is the bacterial density at y0, and λ is the characteristic accumulation length. Measure y0 and b0 from the image and calculate λ through the regression of the distribution by the function.
6. The chemotactic velocity vc is obtained from Eq. (2) using the diffusion coefficient obtained in step B4 (Data analysis) and the accumulation length obtained in step B5 (Data analysis). The chemotactic velocity is measured for an average concentration
c
=
c
1
+
c
2
2
and a gradient
∇
c
=
c
1
-
c
2
L
, where L = 1 mm is the distance between the channels containing the chemoattractant.
7. The chemotactic susceptibility χ(c) is obtained from Eq. (3) by dividing vc by ∇c. The variation of χ(c) with c can be obtained by performing experiments with various couples (c,∇c).
Validation of protocol
We applied this protocol by placing a suspension of E. coli in the central channel between two channels containing 200 and 0 μM of α-methyl-aspartate in the motility buffer. We waited 30 min for the gradient to be established before starting to acquire the films. For this experiment, we then have c = 100 μM and ∇c = 200 μM/mm. Figure 3 shows the first image of three films recorded during the experiment. In the first image, the bacteria, which appear as white dots, are evenly distributed across the image.
Figure 3. Images acquired at three different times (0, 16, and 50 min). The images show the central channels where the bacteria suspension was injected. The observation is done with a 20× objective. The bacteria appear as white dots. Over time, we can see that the bacteria have moved upward and are accumulating on the side closest to the channel containing the chemoattractant.
After a few minutes, the bacteria are more numerous at the top of the image, on the side where the chemoattractant is present. This is shown quantitatively in Figure 4A, where we plotted the profiles b(y) obtained after the image treatment. The adjustment of each profile by an exponential function gives a decay length that we plot as a function of the time at which the films were recorded (Figure 4C). The decay length changes very little over time, indicating that the bacterial concentration profile reaches a stationary regime within a few minutes and no longer evolves over time. The average decay length measured is λ = 130 ± 10 μm (Figure 4C).
Next, each film was processed to determine the velocity correlation function Cy(Δt) (see Figure 4B). Fitting it gives the correlation time τ, which, combined with the average velocity of the bacteria along the y-axis vy, gives the diffusion coefficient
D
=
v
y
2
τ
. The diffusion coefficient as a function of time is shown in Figure 4D. The diffusion coefficient shows little variation over time; we obtained D = 260 ± 30 μm2/s. The accumulation length λ and diffusion coefficient D are then used to determine the chemotactic velocity vc for each film. This is shown as a function of time in Figure 4E. The chemotactic velocity also shows little variation with time; we found: vc =1.9 ± 0.4 μm/s.
The experiment was continued with different concentrations c1 and c2 in the channels, which makes it possible to explore other pairs of parameters (c,∇c) and their influence on the chemotactic velocity. Figure 5 shows, in a log-log representation, the chemotactic susceptibility
χ
(
c
)
=
v
c
∇
c
as a function of c. Each point corresponds to one experiment. Outside the window of c represented in Figure 5, no chemotactic responses were detected. The central part of the data are adjusted by a line indicating that over this region we have
χ
(
c
)
∝
χ
0
c
with χ0 = 490 ± 40 μm2/s/μM.
Figure 4. Data processing to obtain a chemotactic velocity. (A) Bacterial concentration profiles b(y) and (B) velocity correlation function Cy(Δt) measured at three different times during one experiment. The solid lines are exponential fit that determine a) the decay length λ and b) the persistence time τ. (C) Time evolution of the decay length λ. (D) Time evolution of the diffusion
D
=
v
y
2
τ
. (E) Chemotactic velocity vc obtained by Eq. (2) that combines the decay length λ and the diffusion coefficient D. The data are from an experiment with c = 100 μM and ∇c = 200 μM/mm.
Figure 5. Evolution of the chemotactic coefficient χ(c) as a function of the average concentration c. For concentrations below 1 and above 104 μM, no chemotactic response was detected. Solid line:
χ
(
c
)
=
χ
0
c
with χ0 = 490 ± 40 μm2/s/μM.
General notes and troubleshooting
General notes
1. Any remaining bubbles in the channels might cause residual flow that will bias the experiment. Special care is needed to avoid them during the different steps.
2. The settings are for fluorescent strains of bacteria. If the bacteria are not fluorescent, an alternative is to use phase contrast image acquisition. In this case, tracking might require a treatment to eliminate artifacts in the images.
3. The bacteria density profile does not evolve anymore after 5–20 min, depending on the experiment. We recommend waiting for 20 min before starting the image acquisition.
4. The acquisition frequency has to be adjusted so that the typical displacement of a bacteria between two successive images is of the order of 2 pixels. For a characteristic swimming velocity of bacteria of ~10 μm/s, an acquisition frequency of 0.1 s fits this requirement.
5. Films can be acquired at different heights above the agar. We did not observe any effect of this parameter on the estimation of χ(c).
6. If you would like to reuse the PDMS mold, then:
a. Before dismounting the microfluidic cell, flush water into the device.
b. After dismounting, clean the PDMS with ethanol, dry it, and remove the dust on the surface with tape.
c. Finally, stick some tape on the PDMS mold. The mold can then be kept and eventually reused.
7. Swimming velocity or diffusion coefficient measurements are also a good way of identifying batch-to-batch variation in bacterial suspension.
Troubleshooting
Problem 1: The liquid PDMS foams out of the container.
Cause: Bubbles develop in the vacuum chamber and form a foam.
Solution: Break the vacuum regularly.
Problem 2: The agar layer breaks when manipulated.
Cause: The layer is still too warm.
Solution: Make sure the plate is cold enough. It should have some condensation on it.
Problem 3: Flow in the central channel.
Cause 1: Agar delamination on the extrema after 3 h under the microscope.
Solution 1: Wet the agar every hour by pouring pure water drops around the chip.
Cause 2: Motion and vibration of the tubings due to air conditioning.
Solution 2: Put a valve between the microtube containing bacteria and the chip.
Cause 3: Deformation of the PDMS mold due to the tubing.
Solution 3: Do not insert the tubing too deep into the PDMS molding. The molding should be ~1 cm thick so that it can hold the tubing and still remain flat.
Supplementary information
The following supporting information can be downloaded here:
1. File S1. Python Analysis script
Acknowledgments
This work was funded by the CNRS MITI (Mission pour les Initiatives Transverses et Interdisciplinaires) 80|PRIME – 2021 program. We also acknowledge Ahmed et al. [1,5] for the work they conducted, which inspired the current paper.
Competing interests
The authors have no conflict of interest to report.
References
Ahmed, T., Shimizu, T. S. and Stocker, R. (2010). Bacterial Chemotaxis in Linear and Nonlinear Steady Microfluidic Gradients. Nano Lett. 10(9): 3379–3385.
Keller, E. F. and Segel, L. A. (1971). Model for chemotaxis. J Theor Biol. 30(2): 225–234.
Bouvard, J., Douarche, C., Mergaert, P., Auradou, H. and Moisy, F. (2022). Direct measurement of the aerotactic response in a bacterial suspension. Phy Rev E. 106(3): e034404.
Pfeffer, W. (1888). Untersuchungen aus dem botanischen Institut zu Tübingen. (Vol. 2). In: Engelmann, W. 1881–1888
Ahmed, T., Shimizu, T. S. and Stocker, R. (2010). Microfluidics for bacterial chemotaxis. Integr Biol. 2: 604–629.
Stehnach, M., Henshaw, R., Floge, S. and Guasto, J. (2024). Multiplexed Microfluidic Platform for Parallel Bacterial Chemotaxis Assays. Bio Protoc. 14(1352): e5062.
Parkinson, J. S. (1978). Complementation analysis and deletion mapping of Escherichia coli mutants defective in chemotaxis. J Bacteriol. 135(1): 45–53.
Tinevez, J. Y., Perry, N., Schindelin, J., Hoopes, G. M., Reynolds, G. D., Laplantine, E., Bednarek, S. Y., Shorte, S. L. and Eliceiri, K. W. (2017). TrackMate: An open and extensible platform for single-particle tracking. Methods. 115: 80–90.
Lovely, P. S. and Dahlquist, F. (1975). Statistical measures of bacterial motility and chemotaxis. J Theor Biol. 50(2): 477–496.
Article Information
Publication history
Received: Sep 11, 2024
Accepted: Dec 8, 2024
Available online: Dec 31, 2024
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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5,185 | https://bio-protocol.org/en/bpdetail?id=5185&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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This is an In Press version of the protocol that has not yet been assigned to an issue.
Peer-reviewed
A Protocol to Purify Human Mediator Complex From Freestyle 293-F Cells
HT Hui-Chi Tang
KT Kuang-Lei Tsai
TC Ti-Chun Chao
In Press, Available online: Jan 01, 2025
DOI: 10.21769/BioProtoc.5185 Views: 66
Reviewed by: Joana Alexandra Costa ReisRohini Ravindran Nair Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Molecular Cell Oct 2024
Abstract
The Mediator, a multi-subunit protein complex in all eukaryotes, comprises the core mediator (cMED) and the CDK8 kinase module (CKM). As a molecular bridge between transcription factors (TFs) and RNA polymerase II (Pol II), the Mediator plays a critical role in regulating Pol II–dependent transcription. Considering its large size and complex composition, conducting in vitro studies on the Mediator complex is challenging, especially when isolating the intact and homogeneous complex from human cells. Here, we present a method to purify the intact CKM-cMED complex from FreeStyle 293-F cells (293-F cells), which offers advantages for performing large-scale protein purification. To isolate the CKM-bound cMED without the presence of Pol II, FLAG-tagged CDK8, a subunit of the CKM complex, was expressed in 293-F cells for purification, as CKM and Pol II are mutually exclusive in their interaction with cMED. The complex is isolated from nuclear extracts through immunoaffinity purification and further purified by glycerol gradient to enhance its homogeneity. This protocol provides a time- and cost-efficient way to purify the endogenous Mediator complex for structural- and functional-based studies.
Key features
• This protocol describes a method for purifying the endogenous Mediator complex, free of Pol II, from 293-F cells.
• Does not require the use of crosslinkers, offering advantages for structural and functional studies.
Keywords: Mediator CKM CDK8 FreeStyle 293 expression system Protein complex purification Glycerol gradient
Graphical overview
Workflow for isolating the endogenous Mediator complex. The workflow outlines the steps from generating the stable cell line to protein purification.
Background
In eukaryotes, the Mediator complex is an evolutionarily conserved transcriptional coactivator composed of 25 subunits in budding yeast and 30 subunits in humans [1]. The Mediator complex consists of a large core (cMED) with three domains: the head, middle, and tail, along with a dissociable module known as the CDK8 kinase module (CKM). The CKM contains four subunits: cyclin-dependent kinase 8 (CDK8), Cyclin C, MED12, and MED13 [2]. The primary function of cMED is to activate Pol II–dependent transcription by recruiting Pol II to promoters [3]. However, this activity is repressed by the CKM through steric hindrance mediated by MED13 [4,5]. Interestingly, recent studies suggest that CDK8 also plays a positive role in gene transcription, though the mechanisms remain poorly understood. Since the CKM serves as a key entry point for developmental and oncogenic signaling via the Mediator complex [6], isolating the CKM-cMED complex is essential for investigating its mechanisms and functions, providing deeper insights into these processes.
The human CKM-cMED complex consists of 30 subunits, and overexpressing each individually requires considerable effort. As a transcriptional coactivator, the Mediator interacts with multiple binding partners, such as RNA pol II, further complicating the process of isolating pure CKM-cMED complex. Therefore, we developed a protocol to obtain a high-quality and large quantity of CKM-cMED complex.
Previously, we established a stable HEK293 cell line expressing CDK8. However, scaling up proved challenging due to the limitations associated with adherent cells. Then, we selected the FreeStyle 293-F cells suspension cell culture system for its ease of expansion and high protein expression yields. Since CKM and Pol II are mutually exclusive for cMED binding, we overexpressed the CDK8, one of the CKM subunits, with a C-terminal FLAG tag, but not any subunits of cMED, in freestyle 293-F cells to prevent RNA Pol II contamination. The size of the FLAG tag, consisting of eight amino acids, is small and specifically recognized by the antibody conjugated to agarose beads. Additionally, the FLAG tag added to the C-terminus of CDK8 did not compromise the stability of the CKM-cMED complex and still maintained its kinase activity. By establishing stable cell lines through antibiotic selection, we could stably express FLAG-tagged recombinant CDK8 in FreeStyle 293-F cells, enabling the collection of large quantities of cells for protein purification.
For protein purification, we began by isolating nuclear extract to enrich the CKM-cMED complex as it is primarily present in the nucleus. Affinity purification was then performed using anti-FLAG M2 affinity gel to ensure sufficient purity and quality to proceed to the next step. To enhance the homogeneity of the protein complex, we further performed glycerol gradient centrifugation without a cross-linker and successfully obtained the functional endogenous CKM-cMED complex. Because the molecular weight of CKM-cMED is very large, density gradient sedimentation is an effective method to separate the complex from relatively small degradation products, disassembled components, and other impurity proteins present in the FLAG elution. This protocol provides a reliable method for isolating the homogenous endogenous CKM-cMED complex from FLAG-CDK8 expressed cells using FLAG-specific immunoprecipitation and glycerol gradient sedimentation for future investigations of structure-function relationships.
Materials and reagents
Biological materials
1. FreestyleTM 293-F cells (Thermo Fisher, catalog number: R79007), containing 1 × 10 cells suspended in 90% FreeStyleTM 293 expression medium and 10% DMSO, stored in liquid nitrogen
2. Plasmid DNA (pcDNA3.1_CDK8-F)
Reagents
1. Lipofectamine 3000 (Invitrogen, catalog number: L3000001)
2. G418 sulfate (Thermo Fisher, catalog number: 11811023)
3. ANTI-FLAG® M2 affinity gel (Sigma, catalog number: A2220)
4. cOmpleteTM, mini, EDTA-free protease inhibitor cocktail (Sigma, catalog number: 50-161-3323)
5. FreeStyleTM 293 expression medium (prewarm to 37 °C before use)
6. Phosphate buffered saline (PBS), 1×, liquid, pH 7.4 (GenDEPOT, catalog number: P2201-050)
7. HEPES (Merck, catalog number: 391338)
8. Magnesium chloride (Sigma, catalog number: M8266)
9. Potassium chloride (Sigma, catalog number: P3911)
10. Dithiothreitol (DTT) (Fisher Scientific, catalog number: BP172-5)
11. Protease inhibitor cocktail (RPI, catalog number: P50600-1)
12. Ethylenediaminetetraacetic acid (EDTA) (Sigma, catalog number: E6758)
13. NP-40 Surfact-AmpsTM detergent solution (Thermo Fisher, catalog number: 28324)
14. Glycerol (Fisher Scientific, catalog number: BP229-4)
15. Sodium chloride (Sigma, catalog number: 567440)
16. beta-mercaptoethanol (Sigma, catalog number: 444203)
17. DYKDDDDK tag Peptide (FLAG peptide) (APExBio, catalog number: A6002)
Solutions
1. Hypotonic buffer (see Recipes)
2. Nuclear extraction buffer (see Recipes)
3. Washing buffer A (see Recipes)
4. Washing buffer B (see Recipes)
5. Washing buffer C (see Recipes)
6. Elution buffer (see Recipes)
7. 15% glycerol gradient buffer (see Recipes)
8. 40% glycerol gradient buffer (see Recipes)
Recipes
1. Hypotonic buffer
10 mM HEPES pH 7.9
1.5 mM MgCl2
10 mM KCl
0.5 mM dithiothreitol (DTT)
Protease inhibitors (freshly added)
2. Nuclear extraction buffer
25 mM HEPES pH 7.9
1.5mM MgCl2
0.6 M KCl
0.5 mM DTT
0.2 mM EDTA
0.02% NP-40
20% glycerol
Protease inhibitors (freshly added)
3. Washing buffer A
25 mM HEPES pH 7.6
1.5 mM MgCl2
150 mM NaCl
10% glycerol
0.2 mM EDTA
2 mM beta-mercaptoethanol
Protease inhibitors (freshly added)
4. Washing buffer B
25 mM HEPES pH 7.6
1.5 mM MgCl2
150 mM NaCl
10% glycerol
0.2 mM EDTA
2 mM beta-mercaptoethanol
5. Washing buffer C
25 mM HEPES pH 7.6
1.5 mM MgCl2
150 mM NaCl
10% glycerol
0.2 mM EDTA
0.01% NP40
2 mM beta-mercaptoethanol
6. Elution buffer
25 mM HEPES pH 7.6
1.5 mM MgCl2
150 mM NaCl
10% glycerol
0.2 mM EDTA
0.01% NP40
2 mM beta-mercaptoethanol
200 μg/mL 1× FLAG peptide
7. 15% glycerol gradient buffer
20 mM HEPES pH 7.6
1 mM MgCl2
300 mM NaCl
0.25 mM EDTA
1 mM beta-mercaptoethanol
15% (v/v) glycerol
8. 40% glycerol gradient buffer
20 mM HEPES pH 7.6
1 mM MgCl2
300 mM NaCl
0.25 mM EDTA
1 mM beta-mercaptoethanol
40% (v/v) glycerol
Filter sterilize and store all buffers at 4 °C.
Laboratory supplies
1. Clean, sterile glass 250, 1,000, 2,000 mL flasks
2. Hemacytometer
3. 60 × 15 mm non-treated culture dish, sterile (CytoOne, catalog number: CC7672-3359)
4. 15 and 50 mL conical sterile polypropylene centrifuge tubes (Thermo Scientific, catalog number: 12-565-269, 12-565-271)
5. Dounce homogenizer
6. 1.7 mL microtubes (e.g., Axygen, catalog number: 14-222-168)
7. Centrifuge bottles with sealing closure (Thermo Scientific, catalog number: 05-564-3)
8. Polycarbonate ultracentrifuge bottles (Beckman, catalog number: 355622)
9. 4 mL ultra-clear polypropylene tubes (Beckman, catalog number: 344062)
10. Econo-Pac® chromatography columns (Bio-Rad, catalog number: 7321010)
11. NovexTM tris-glycine mini protein gels, 4%–20%, 1.0 mm, WedgeWellTM format (Invitrogen, catalog number: XP04205BOX)
Equipment
1. Standard orbital shaker (VWR, model: 1000, catalog number: 89032-088)
2. CO2 cell culture incubator
3. Inverted microscope
4. Biological safety cell culture hood
5. 37 °C water bath
6. Benchtop centrifuge (Eppendorf, catalog number: 022625501)
7. High-speed centrifuge (Thermo Sorvall, model: RC-6 Plus; PTI FiberLite F10-6x500y fixed angle rotor)
8. Ultracentrifuges (Beckman, model: Optima XPN-90; 45 Ti fixed-angle titanium rotor, SW60 Ti swinging-bucket rotor)
9. GRADIENT MASTERTM (Biocomp, Model 108)
10. Tube revolver rotator (Thermo Scientific, catalog number: 88881001)
Software and datasets
1. BioRender (https://www.biorender.com/) was used for the Graphical overview: Tang, H. (2025) https://BioRender.com/z21d813; and Figure 3: Tang, H. (2025) https://BioRender.com/t82q193
Procedure
A. Overexpression of CDK8-FLAG in FreeStyleTM 293-F cells
1. Cell culture
a. Thaw the FreeStyleTM 293-F cells quickly in a 37 °C water bath.
b. Before the vial is almost thawed, clean the vial with 70% ethanol.
c. Gently transfer the cells from the vial to the 250 mL flask containing 30 mL of prewarmed FreeStyleTM 293 expression medium inside a biosafety cabinet (cell density is around 0.33 × 106 cells/mL).
d. Place the flask on the orbital shaker (at 135 rpm) inside the incubator at 37 °C with a humidified atmosphere of 8% CO2.
e. Maintain the cells at 0.3–3 × 106 cells/mL density in a 250 mL flask.
Note: Cells should be healthy with viability greater than 90%. Low viability leads to reduced transfection efficiency.
2. Transfection and generation of stable cell line
Day 0
a. Culture 293-F cells at a density of 0.6 × 106 to 0.7 × 106 cells/mL.
Day 1
a. Measure the cell density using a hemacytometer.
b. Dilute the cells to 1 × 106 cells/mL.
c. Seed the cells in a volume of 3 mL in the 6 cm non-treated plate.
d. Dilute 3 μg of plasmid DNA and 6 μL of P3000 into 100 μL of FreeStyleTM 293 expression medium in a 1.7 mL tube.
e. Dilute 7.5 μL of lipofectamine into 100 μL of FreeStyleTM 293 expression medium in another tube.
f. Gently add the diluted lipofectamine (from step A, day 1, e) into the diluted DNA solution (from step A, day 1, d).
g. Incubate the mixture for 10–15 min at room temperature.
h. Drop the DNA–lipid complex into cells.
i. Place the 6 cm non-treated plate on the orbital shaker inside the incubator at 37 °C with 8% CO2.
Day 2
a. Change the fresh media after 8–12 h.
b. Collect the cells in a 15 mL tube.
c. Centrifuge the cells at 201× g for 3 min.
d. Discard the supernatant and resuspend the cells in 4 mL of prewarmed fresh medium.
e. Transfer the cells into the 6 cm non-treated plate.
Day 4
a. Add G418 sulfate to a final concentration of 200 μg/mL for selection.
b. Measure the cell density and viability using a hemacytometer. Change the fresh media every 3–4 days.
c. When cell density is higher than 1 × 106 cells/mL, split the cells into a 1:4 ratio in fresh medium and use a smaller amount of G418 (50 μg/mL) for maintenance.
d. Collect a small amount of cells and check protein expression using western blot (Figure 1). This is an essential checkpoint for the following steps.
e. Expand the cells to 4 L (check protein expression frequently using western blot while expanding cells).
f. Pour the medium into centrifuge bottles from glass flasks. Harvest the cells by centrifugation at 1,590× g for 10 min at 4 °C in a Thermo Sorvall RC-6 Plus Centrifuge with F10-6x500y fixed angle rotor.
g. Discard the supernatant and resuspend the cell pellet with ice-cold PBS.
h. Transfer the cells into 50 mL tubes.
i. Centrifuge at 201× g for 10 min at 4 °C.
j. Discard the supernatant and store the pellet at -80 °C.
Figure 1. Expression test of hCDK8-FLAG. FreeStyleTM 293-F cells were collected after transfection. FLAG antibody was used for western blot analysis. C: FreeStyleTM 293-F cells; Lane 1: hCDK8-FLAG overexpressed in FreeStyleTM 293-F cells. Sample was collected when expanding the cell culture.
B. Purification of human cMED-CKM
1. Cell lysis and nuclear extraction
a. Defrost 40 g of 293F cell pellets stably over-expressing FLAG-CDK8 in 250 mL of pre-chilled hypotonic buffer.
b. Resuspend the cell pellet by gently pipetting.
c. Incubate on ice for 10 min.
d. Use a Dounce homogenizer (30 mL) with 20 gentle strokes to lyse the cells.
e. Aliquot the lysate into four ultracentrifuge bottles for ultracentrifugation.
f. Centrifuge (45 Ti Rotor) at 46,377.8× g for 30 min at 4 °C and discard the supernatant.
g. Resuspend the nuclear pellet with 40 mL/tube of hypotonic buffer.
h. Centrifuge (45 Ti Rotor) at 46,377.8× g for 10 min at 4 °C and discard the supernatant.
i. Resuspend the nuclear pellet with 40 mL/tube of hypotonic buffer.
j. Centrifuge (45 Ti Rotor) at 46,377.8× g for 10 min at 4 °C and discard the supernatant.
k. Resuspend all nuclear pellets with 25 mL of nuclease extraction buffer.
l. Add hypotonic buffer to the final volume of 50 mL (final salt concentration: 300 mM KCl).
m. Transfer the suspension from the 50 mL tube into the polycarbonate ultracentrifuge bottles.
n. Rotate the suspension for 1 h at 4 °C.
o. Centrifuge (45 Ti Rotor) at 126,263.7× g for 1 h at 4 °C.
2. Binding
a. Collect the supernatant and aliquot into a 50 mL tube.
b. Get 4 mL of ANTI-FLAG® M2 affinity gel in another 15 mL tube.
c. Equilibrate the resin with 10 mL of pre-chilled nuclear extraction buffer, centrifuge at 201× g for 1 min, and discard the supernatant.
d. Repeat step B2c twice.
e. Apply the resin to the nuclear extract from step B2a and incubate the sample for 4 h at 4 °C with end-over-end rotation. The incubation time is associated with the purification yield.
3. Wash
a. Load the mixture of resin and the nuclear extract into the column.
b. Wash the resin with 30 mL of washing buffer A.
c. Wash the resin with 30 mL of washing buffer B.
d. Wash the resin with 30 mL of washing buffer C.
4. cMED-CKM complex elution
a. Seal the bottom of the column and add 8 mL of elution buffer from the top.
b. Seal the top of the column and incubate the sample for 1 h at 4 °C with end-over-end rotation.
c. Collect the elution (8 mL) and run SDS-PAGE to perform a silver stain to confirm purity (Figure 2). This is a critical checkpoint for assessing the purity of the elution before further purification.
d. Concentrate the eluted solution to approximately 2.4 mL.
Figure 2. FLAG purification result of the Mediator complex, 4%–20% gradient gel, silver stained. The elution fraction was collected after FLAG purification. The CKM subunits, including MED12 and MED13, are indicated by green text.
5. Glycerol gradient
a. Add 1,860 μL of 40% glycerol gradient buffer (heavy solution) into ultra-clear polypropylene tubes. Using P1000 Pipetman, load 930 μL of the buffer into the tube twice.
b. Add 1,860 μL of 15% glycerol gradient buffer (light solution) gently into the same tube. Using P1000 Pipetman, load 930 μL of the buffer into the tube twice.
c. Avoid air bubble formation when adding buffer or samples. This will disturb the layers.
d. Make sure the heavy–light interface rises precisely to the middle of the tube and seal the tube.
e. Prepare six tubes. The detailed procedure is described in Figure 3.
Figure 3. Flowchart of glycerol gradient
f. Use the short rubber cap to seal the tube and make sure the side with the hole is the last part to seal.
g. Place the tubes in the tube holder and use the Gradient Master instrument to rotate and mix the solutions to form the gradient. Choose the gradient 15%–40% and hit Run.
h. Remove the seal and load 400 μL of cMED-CKM complex elution by gently pipetting on top of the gradients.
i. Balance the tubes before placing them into the rotor. Caution: an improperly loaded rotor or weight difference will lead to rotor imbalance.
j. Centrifuge (SW60 Ti Rotor) at 120,937.6× g for 17 h at 4 °C.
k. Fractionate the gradients by collecting 200 μL of sample per fraction at 4 °C and analyze the fraction using SDS-PAGE (Figure 4A) and western blot (Figure 4B).
Figure 4. Overview of the glycerol gradient results. The eluted samples obtained by anti-FLAG affinity gel from Cdk8-FLAG-expressed 293-F cells were applied to glycerol gradient sedimentation. Fractions were collected from top to bottom and analyzed using a 4%–20% SDS-PAGE gel for silver stained (A) and immunoblot (B). FLAG and MED6 antibodies were used. M: Marker; 1. Fraction 1–3; 2. Fraction 4–6; 3. Fraction 7–8; 4. Fraction 9–10; 5. Fraction 11–12; 6. Fraction 13–14; 7. Fraction 15–16; 8. Fraction 17–18; 9. Fraction 19–21.
Validation of protocol
This protocol has been used in the following research article:
• Chao et al. [7]. Structural basis of the human transcriptional Mediator regulated by its dissociable kinase module. Mol Cell. https://doi.org/10.1016/j.molcel.2024.09.001 (Figure 1B, Figure S1.B). The cryo-EM structure of cMED-CKM complex were obtained followed the purification steps from this protocol.
General notes and troubleshooting
General notes
1. In this protocol, we use 293-F cells to overexpress the CDK8-F as an example to isolate the mediator complex. It also applies to other mammalian expression strains (e.g., HEK293 cells).
2. The protocol is adaptable for expression at any scale. From 1 L of 293-F cell culture, approximately 10 g of pellet can be expected.
3. Recommended working volume for the flasks: 600 mL of cells for a 2 L shake flask and 50 mL of cells for a 250 mL flask.
Troubleshooting
A. Low transfection efficiency
1. Cell viability maintained at >90% is recommended. Use fresh cell stocks or early-passage cells.
2. Check the quality of the plasmid DNA. (Prepare the plasmid DNA with low endotoxin and contamination.)
3. Vary the amount of DNA or transfection reagent. Try 3–6 μg of plasmid DNA or 3.75–7.5 μL of lipofectamine reagent.
4. Incubation time of the mixture should be between 10 and 20 min. Longer times can lead to loss of transfection efficiency.
B. Low protein yield
1. Check the protein expression using western blot for each passage. Prolonged culture (>30 passages) may reduce the protein yield. Thaw a new batch if necessary.
2. Check the cell lysis efficiency using SDS-PAGE/western blot analysis. Increase the strokes when lysing the cells if necessary.
3. Increase the incubation time for the cell lysate with ANTI-FLAG® M2 affinity gel.
4. Verify the concentration of the FLAG peptide in the elution buffer to reach a successful protein elution.
C. Glycerol gradient shift
1. Verify the glycerol concentration using a refractometer.
2. Carefully load the gradient buffer without disturbance or air bubbles formation.
3. Ensure that a clear interface forms between the two layers before placing the tubes in the tube holder. Hold the tube steady and do not disturb the interface.
4. After adding the FLAG elution on top of the gradient, confirm a distinct interface between the FLAG elution and the gradient.
D. Poor protein purity
1. Increase the extra wash steps (e.g., 5 washes) to reduce non-specific binding if necessary.
2. Keep the samples on ice to avoid protein aggregation or degradation.
Acknowledgments
The protocol presented here is adapted from Chao et al. [7]. This work was supported by US National Institutes of Health grant R01 GM143587 (K.L.T.).
Competing interests
The authors declare no competing interests.
References
Verger, A., Monte, D. and Villeret, V. (2019). Twenty years of Mediator complex structural studies. Biochem Soc Trans. 47(1): 399–410.
Tsai, K. L., Tomomori-Sato, C., Sato, S., Conaway, R. C., Conaway, J. W. and Asturias, F. J. (2014). Subunit Architecture and Functional Modular Rearrangements of the Transcriptional Mediator Complex. Cell. 158(2): 463.
Soutourina, J. (2018). Transcription regulation by the Mediator complex. Nat Rev Mol Cell Biol. 19(4): 262–274.
Tsai, K. L., Sato, S., Tomomori-Sato, C., Conaway, R. C., Conaway, J. W. and Asturias, F. J. (2013). A conserved Mediator-CDK8 kinase module association regulates Mediator-RNA polymerase II interaction. Nat Struct Mol Biol 20(5): 611–619.
Elmlund, H., Baraznenok, V., Lindahl, M., Samuelsen, C. O., Koeck, P. J., Holmberg, S., Hebert, H. and Gustafsson, C. M. (2006). The cyclin-dependent kinase 8 module sterically blocks Mediator interactions with RNA polymerase II. Proc Natl Acad Sci USA. 103(43): 15788–15793.
Clark, A. D., Oldenbroek, M. and Boyer, T. G. (2015). Mediator kinase module and human tumorigenesis. Crit Rev Biochem Mol Biol. 50(5): 393–426.
Chao, T. C., Chen, S. F., Kim, H. J., Tang, H. C., Tseng, H. C., Xu, A., Palao, L., Khadka, S., Li, T., Huang, M. F., et al. (2024). Structural basis of the human transcriptional Mediator regulated by its dissociable kinase module. Mol Cell. 84(20): 3932–3949.e10.
Article Information
Publication history
Received: Oct 3, 2024
Accepted: Dec 3, 2024
Available online: Jan 1, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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5,186 | https://bio-protocol.org/en/bpdetail?id=5186&type=0 | # Bio-Protocol Content
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This is an In Press version of the protocol that has not yet been assigned to an issue.
Peer-reviewed
Closed Systems to Study Plant–Filamentous Fungi Associations: Emphasis on Microscopic Analyses
VS Vasiliki Skiada
KP Kalliope K. Papadopoulou
In Press, Available online: Jan 05, 2025
DOI: 10.21769/BioProtoc.5186 Views: 103
Reviewed by: Shweta Panchal Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Fungal Genetics and Biology Jun 2019
Abstract
In nature, filamentous fungi interact with plants. These fungi are characterized by rapid growth in numerous substrates and under minimal nutrient requirements. Investigating the interaction of these fungi with their plant hosts under controlled conditions is of importance for many researchers aiming to proceed with molecular or microscopical investigations of their favorite plant–fungus interaction system. The speed of growth of these fungi complicates transferring plant–fungal interaction systems in laboratory conditions. The issue is more complicated when monoxenic conditions are desired, to ensure that only two members (a fungus and a plant) are present in the system under study. Here, two simple closed systems for investigating plant–filamentous fungi associations under laboratory, monoxenic conditions are described, along with their limitations. The plant and fungal growth conditions, methods for sampling, staining, sectioning, and subsequent microscopical imaging of colonized plant tissues with affordable, common laboratory tools are described.
Key features
• Setting up closed systems for microscopical observations of plant–filamentous fungi (emphasis on model legumes–Fusaria) associations and temporal in vivo observations of the association(s).
• Preparing root samples for microscopical observations: staining, sectioning, and mounting on microscopical slides.
• Using low-cost equipment for performing microscopical observations and imaging.
• Using fluorescence microscopy: provision of common fluorophores to highlight specific plant and fungal tissues, compartments, and structures.
Keywords: Plant–microbe interactions Filamentous fungi Lotus japonicus Fusarium solani Fusarium oxysporum Axenic system Microscopy Staining techniques Tissue sectioning Fluorochromes
Graphical overview
Graphical overview of closed systems used to study plant–filamentous fungi associations and subsequent tissue processing for microscopic analyses. Lotus japonicus seeds are scarified, decontaminated, and germinated in Petri dishes. A Fusarium strain of choice is grown to collect propagules for plant inoculations. L. japonicus plantlets are transplanted in magenta boxes/glass jars or in square Petri dishes and inoculated with the fungus. Square Petri dishes are recommended for in vivo observations. For further tissue(s) processing, root/stem/leaf tissues are collected, embedded in agarose or in paraffin wax, and sectioned by hand, in cases where sectioning is necessary. Either whole root/stem/leaf tissues or tissue sections are stained to highlight inter- and/or intracellular fungal structures. Stained tissues are mounted on slides and observed using a compound microscope. An epifluorescence microscope is used when tissues are stained with fluorescent dyes to highlight specific plant tissues and compartments, fungal cells, and structures. A confocal system is used, if accessible, for sharper images and/or for live cell imaging.
Background
Plant–microbe interactions (PMIs) are of fundamental importance and occur everywhere in nature. Multiple microbial partners interact with root tissues and some of them will simultaneously colonize plant tissues. PMI research greatly benefits from the ability to grow plants in axenic (sterile) conditions [1]. Even in simplified systems, difficulties arise in distinguishing the partner responsible for a given response. Investigating individual, simplified, bipartite associations of the plant with a single microbial partner aids in understanding the reciprocal impact and deciphering specific mechanisms of response. Transferring PMIs in controlled laboratory conditions is of importance, especially when basic biological questions either related to molecular or cellular biology are to be answered. To perform and investigate PMIs under gnotobiotic conditions, model systems are necessary. The construction of model systems where plant roots associate with one or more selected microbial species allows researchers to investigate interactions in simpler systems than those occurring in nature [2].
In monoxenic conditions, for example, a plant species associates with only one microbial partner. Several closed systems have been reported, including agar plates, which have been used extensively in mycorrhizal research [3]. Here, we focus on the interaction of model plants with filamentous fungi. When investigating plant–filamentous fungi associations at the molecular and cellular level, setting up a model, closed system where the interaction will occur is important. The adjustment of several parameters that depend on the organism under investigation and the research questions posed should be considered. In any given model system, further processing of infected plant tissues depends on the research aim. For many studies, determining the degree of root colonization by fungi is necessary, as correlations to benefits for the plants may be inferred, such as promotion of growth and seed yield as well as tolerance against biotic and abiotic stress, or even correlations to loss of benefits, in cases of fungal over-colonization. In this respect, powerful staining techniques are required to visualize and estimate fungal growth within the roots [4].
Sectioning of plant tissues is also considered when a more detailed analysis is required, such as in determining the exact location of fungal cells or the infected plant cells and tissues, as well as revealing alterations on the plant/fungal side. Besides conventional microscopic analysis, the use of fluorescent dyes is also sometimes necessary to highlight specific colorless plant or fungal compartments, tissues, and/or deposition of plant substances, such as lignin and suberin [5–7].
Here, we describe in detail two closed, monoxenic systems of model legume plants with filamentous fungi by taking advantage of the previously described model interaction system of the legume Lotus japonicus with the endophytic, beneficial fungal strain Fusarium solani strain K (FsK) [8–10]. We emphasize the use of affordable tools and materials for performing these experiments. Furthermore, we describe the processing of tissues, including harvest, staining, and sectioning to visualize inter- and intracellular fungal structures to perform basic microscopic analysis. Finally, we present common fluorophores that one may consider if a more detailed microscopic analysis, using fluorescence microscopy, is required.
Materials and reagents
Biological materials
1. L. japonicus seeds (ecotype Gifu)
2. Fusarium strain [such as F. solani strain K (FsK), F. oxysporum forma specialis (f. sp.) medicaginis (Fom), Fusarium oxysporum f. sp. lycopersici (Fol); and Fusarium oxysporum f. sp. radicis-lycopersici (Forl) (see General note 1)]
Reagents
1. Deionized water
2. Sulfuric acid (H2SO4) (Sigma-Aldrich, catalog number: 258105)
3. Potato Dextrose Broth (Condalab, catalog number: 1261)
4. Commercial bleach (non-concentrated, containing ~5% w/v sodium hypochlorite, generic)
5. Murashige and Skoog (MS) medium including vitamins (Duchefa Biochemie, catalog number: M022)
6. Bacteriological Agar (Condalab, catalog number: 1800)
7. Sodium chloride (NaCl) (Sigma-Aldrich, catalog number: S9888)
8. Potassium nitrate (KNO3) (Chem-Lab NV, catalog number: CL00.1164)
9. Calcium nitrate tetrahydrate [Ca(NO3)2·4H2O] (Chem-Lab NV, catalog number: CL00.0340)
10. Potassium dihydrogen phosphate (KH2PO4) (Chem-Lab NV, catalog number: CL00.1146)
11. Magnesium sulfate heptahydrate (MgSO4·7H2O) (Chem-Lab NV, catalog number: CL00.1324)
12. Boric acid (H3BO3) (Chem-Lab NV, catalog number: CL00.0216)
13. Manganese(II) chloride tetrahydrate (MnCl2·4H2O) (Chem-Lab NV, catalog number: CL00.1328)
14. Zinc sulfate heptahydrate (ZnSO4·7H2O) (Chem-Lab NV, catalog number: CL00.2629)
15. Copper sulfate pentahydrate (CuSO4·5H2O) (Chem-Lab NV, catalog number: CL00.1127)
16. Molybdic acid (MoO3·H2O) (Supelco, catalog number: 100400)
17. Potassium chloride (KCl) (Chem-Lab NV, catalog number: CL00.1133)
18. Sodium ferric ethylenediaminetetraacetic acid (NaFeEDTA) (Sigma-Aldrich, catalog number: E6760)
19. Potassium iodide (KI) (Chem-Lab, catalog number: CL00.1134)
20. Sodium molybdate dihydrate (Na2MoO4·2H2O) (Applichem, catalog number: 131701)
21. Glycine (Duchefa Biochemie, catalog number: G0709)
22. Thiamine hydrochloride (Duchefa Biochemie, catalog number: T0614)
23. Pyridoxine Hydrochloride (Duchefa Biochemie, catalog number: P0612)
24. Nicotinic acid (Duchefa Biochemie, catalog number: N0611)
25. Myo inositol (Duchefa Biochemie, catalog number: I0609)
26. Agarose (low melt, Carl Roth, catalog number: 6351)
27. Ethanol (PanReac Applichem, catalog number: 147194)
28. Potassium hydroxide (KOH) (pellets, Chem-Lab, catalog number: CL00.1135)
29. Hydrochloric acid (HCl) (37+% HCl, Chem-Lab, catalog number: CL00.0310)
30. Chlorazol black (CB) (Sigma-Aldrich, catalog number: C1144)
31. Glycerol (PanReac Applichem, catalog number: 131339)
32. Lactic acid (PanReac Applichem, catalog number: 141034)
33. Trypan blue (TB) (Sigma-Aldrich, catalog number: T6146)
34. Black ink (Sheaffer, catalog number: 94231)
35. Acetic acid (BioChemica, catalog number: 131008)
36. Polyvinyl acetate (PVA) (Carl Roth, catalog number: 9154)
37. Liquid nitrogen (to collect tissues for subsequent molecular analysis, e.g., gene expression) (generic)
38. Paraffin wax flakes (generic)
Solutions
1. Potato dextrose broth medium (PDB) (prepared as per purchased product instructions)
2. Commercial bleach (containing ~5% w/v sodium hypochlorite) solution (20% v/v) (see Recipes)
3. Half-strength Murashige and Skoog (MS) medium including vitamins (see Recipes)
4. Isotonic solution (NaCl 0.85% w/v) (see Recipes)
5. Hoagland medium (see Recipes)
6. M medium (see Recipes)
7. Low melting temperature agarose solution (0.4% w/v) (see Recipes)
8. Ethanol solution (70% v/v) (see Recipes)
9. KOH (10% w/v) (see Recipes)
10. HCl (1 N) (see Recipes)
11. CB stain solution (0.05% w/v in water:glycerol:lactic acid 1:1:1) (see Recipes)
12. TB stain solution (0.05% w/v in water:glycerol:lactic acid 1:1:1) (see recipes)
13. Black ink (5% v/v in 5% v/v acetic acid) (see Recipes)
14. Agarose solution (6%–8% w/v) (see Recipes)
15. Polyvinyl-Lacto-Glycerol (PVLG) (see Recipes)
16. Glycerol (50%–90% v/v) (see Recipes)
Recipes
1. Commercial bleach solution (20% v/v)
Reagent Final concentration Quantity or Volume
Commercial bleach (containing ~5% w/v sodium hypochlorite) 20% 10 mL
H2O n/a 40 mL
Total n/a 50 mL
2. Half-strength MS medium including vitamins [11]
Reagent Final concentration Quantity or Volume
MS medium 0.5× 2.2 g
Agar 1% 10 g
Total n/a 1,000 mL
Adjust the pH of the medium to 5.5 before sterilization. Sterilize at 121 °C for 15 min.
3. Isotonic solution (NaCl 0.85% w/v)
Reagent Final concentration Quantity or Volume
NaCl 0.85% 0.85 g
Total n/a 100 mL
Sterilize at 121 °C for 15 min.
4. Hoagland medium [12,13]
Reagent Final concentration Quantity or Volume
KNO3 5 mM 5 mL of 1 M
Ca(NO3)2·4H2O 5 mM 5 mL of 1 M
KH2PO4 1 mM 1 mL of 1 M
MgSO4·7H2O 2 mM 2 mL of 1 M
FeEDTA 1–5 mg/L 1–5 mL of 1,000 mg/L
Micronutrient stock solution* - 1 mL
Total n/a 1,000 mL
*Micronutrient stock solution for Hoagland medium
Reagent Final concentration Quantity or Volume
H3BO3 - 2.86 g
MnCl2·4H2O - 1.81 g
ZnSO4·7H2O - 0.22 g
CuSO4·5H2O - 0.08 g
MoO3·H2O - 0.02 g
Total n/a 1,000 mL
If necessary, sterilize the medium at 121 °C for 15 min.
5. M medium [14]
Reagent Final concentration Quantity or Volume
MgSO4·7H2O - 731 mg
KNO3 - 80 mg
KCl - 65 mg
KH2PO4 - 4.8 mg
Ca(NO3)2·4H2O - 288 mg
NaFeEDTA - 8 mg
KI - 0.75 mg
MnCl2·4H2O - 6 mg
ZnSO4·7H2O - 2.65 mg
H3BO3 - 1.5 mg
CuSO4·5H2O - 0.13 mg
Na2MoO4·2H2O - 0.0024 mg
Glycine - 3 mg
Thiamine hydrochloride - 0.1 mg
Pyridoxine Hydrochloride - 0.1 mg
Nicotinic acid - 0.5 mg
Myo inositol - 50 mg
Agar 0.4%–0.8% 4–8 g
Total n/a 1,000 mL
Adjust the pH of the medium to 5.5 before sterilization. Sterilize at 121 °C for 15 min.
6. Low melting temperature agarose solution (0.4% w/v)
Reagent Final concentration Quantity or Volume
Agarose 0.4% 0.1 g
Total n/a 25 mL
7. Ethanol solution (70% v/v)
Reagent Final concentration Quantity or Volume
EtOH 70% 70 mL
Total n/a 100 mL
8. KOH (10% w/v)
Reagent Final concentration Quantity or Volume
KOH 10% 10 g
Total n/a 100 mL
9. HCl (1 N)
Reagent Final concentration Quantity or Volume
HCl (37% w/v, 12.08 N) 1 N 8.28 mL
Total n/a 100 mL
10. CB stain solution (0.05% w/v in water:glycerol:lactic acid 1:1:1)
Reagent Final concentration Quantity or Volume
Chlorazol black 0.05% 0.075 g
H2O - 50 mL
Glycerol - 50 mL
Lactic acid - 50 mL
Total n/a 150 mL
11. TB stain solution (0.05% w/v in water:glycerol:lactic acid 1:1:1)
Reagent Final concentration Quantity or Volume
Trypan blue 0.05% 0.075 g
H2O - 50 mL
Glycerol - 50 mL
Lactic acid - 50 mL
Total n/a 150 mL
12. Black ink stain solution (5% v/v in 5% v/v acetic acid) [15]
Reagent Final concentration Quantity or Volume
Black ink (Sheaffer) 5% 5 mL
Acetic acid 5% 5 mL
H2O - 90 mL
Total n/a 100 mL
13. Agarose solution (6%–8% w/v)
Reagent Final concentration Quantity or Volume
Agarose 6%–8% 6–8 g
Total n/a 100 mL
14. PVLG
Reagent Final concentration Quantity or Volume
Water - 100 mL
Lactic acid - 100 mL
Glycerol - 10 mL
PVA - 16.6 g
Total n/a 210 mL
15. Glycerol (50%–90% v/v)
Reagent Final concentration Quantity or Volume
Glycerol 50%–90% 15–27 mL
Total n/a 30 mL
Laboratory supplies
1. Eppendorf tubes (1.5 and 2.0 mL) (D. Dutscher, catalog numbers: 033511 and 033297)
2. Glass Pasteur pipette(s) (D. Dutscher, catalog number: 251564)
3. Rubber teat (pipetting aid for Pasteur pipettes) (Sigma-Aldrich, catalog number: Z111597)
4. Pipette tips (Kisker, catalog numbers: VL700G and VL004G for 1-200 and 100-1000μl, respectively)
5. Pipette(s) (Gilson, catalog number: F144058M and F144059M for Pipetman P200 and P1000, respectively)
6. Forceps (generic)
7. Petri dishes (94/16 mm, Greiner Bio-One, catalog number: 633181)
8. Square Petri dishes (120/120/17 mm, Greiner Bio-One, catalog number: 688102)
9. Aluminum foil (generic)
10. Scalpel (generic)
11. Parafilm (D. Dutscher, catalog number: 090260)
12. Sterile cheesecloth (generic)
13. Sterile glass funnel (Boro Simax)
14. Sterile Falcon tubes (Sarstedt, catalog number: 62.547.254)
15. Neubauer chamber (hemacytometer) (Marienfeld Superior, catalog number: 0640030)
16. Cover glass for Neubauer chamber (20x26mm, Marienfeld Superior, catalog number: 0350000)
17. Magenta boxes (Phyta Jar, 74 × 71 × 98 mm, HIMEDIA, catalog number: PW1138-50NO), or
18. Glass jars with lid (~370 mL) (generic)
19. Substrate (sand:vermiculite mixture, 2:1) (generic)
20. Injection syringe & needle (generic)
21. Dissecting (inoculation) needle (generic)
22. Gas permeable plastic film (25-μm) (Film 25, Lumox, Sarstedt, catalog number: 94.6077.316)
23. Filter paper (generic)
24. Small plastic zip bags, Falcon tubes, or similar item(s) for tissue storage
25. Small strainer (generic)
26. Razor blades (generic)
27. Plastic Pasteur pipette (with cut end) (Biosigma, catalog number: BSV136)
28. Conical flask or beaker (Boro Simax)
29. Silicone rubber flat embedding mold (Agar Scientific, AGG3530), or
30. Molds made of aluminum foil
31. Microscope slides (Knittel, 76x26mm, catalog number: 303160)
32. Coverslips (Knittel, 24x50mm, catalog number: VD12450Y1A.01)
33. Nut (generic)
34. Bolt (~2 cm in depth) (generic)
35. Fine point forceps, or brush (generic)
36. Gloves (generic)
37. Lab coat (generic)
Equipment
1. Tube vortex mixer (Velp Scientifica)
2. Fume hood (Equip LaboTM Fume Hood)
3. Laminar flow hood (Telstar, Bio II Advance Plus 4)
4. Shaking incubator (Labtech)
5. Autoclave (Raypa, AES-50)
6. Refrigerator (4 °C) (generic)
7. Plant growth chamber (grow tent for indoor cultivation of plants, custom-made)
8. Centrifuge (Hermle)
9. Water bath (JP Selecta)
10. Hot plate stirrer (for agarose melting) (Agimatic-E)
11. Stereoscope (Novex Holland)
12. Oven (JP Selecta)
13. Compound light microscope (Novex Holland)
14. Fluorescence microscope (Olympus BX53 upright microscope)
15. Confocal microscope system (Zeiss LSM800 system)
16. Camera for imaging (Olympus DP74)
Software and datasets
1. Appropriate software for image acquisition (here, the cellSens imaging software was used)
Procedure
A. Seed scarification, decontamination, and germination
1. Scarify and decontaminate L. japonicus seeds: Briefly, chemically scarify L. japonicus seeds with a ~12–20 min sulfuric acid treatment (by shaking the Eppendorf tube containing the seeds and sulfuric acid on a tube vortex mixer), wash thoroughly with deionized water, surface sterilize seeds for 20 min in a 20% v/v NaOCl solution, and wash 6× with sterile deionized water (see General note 2).
2. Keep seeds in water at 4 °C overnight in an Eppendorf tube covered with aluminum foil to promote seed germination. Afterward, place them in Petri dishes containing half-strength (1/2) MS medium [11] and transfer the Petri dishes to a growth chamber with a 16/8 h light/dark photoperiod at 22 °C. In the following experimental procedures, 6–12-days-old L. japonicus plantlets are used.
B. Fungal inoculum preparation
1. Perform routine culture of the selected Fusarium strain by inoculating a fresh potato dextrose agar plate with a small fungal plug from a previous fungal culture on the same medium.
2. Isolate fungal asexual reproductive spores (conidia) as follows. Inoculate a flask containing potato dextrose broth (100 mL) with a fungal plug from the periphery of the fungal colony of the routine culture of step 1. Allow the culture to grow for ~5 days (26–28 °C, 160 rpm). Filter the culture through a sterile cheesecloth (using a sterile glass funnel) to separate the fungal mycelium from fungal conidia. Centrifuge the culture filtrate containing the fungal conidia (4,000× g), remove the supernatant, and resuspend the recovered conidia in a few milliliters of isotonic solution (0.85% w/v NaCl).
3. Dilute the suspension of fungal conidia appropriately and determine conidia concentration by using a Neubauer chamber and a compound light microscope [8] (see General note 3).
Note: For FsK, conidia collected are initially diluted in ~5 mL of isotonic solution, and conidia concentration is determined in this solution using a Neubauer chamber. Conidia are further diluted to a working concentration of 100 conidia per 200 μL of isotonic solution (i.e., 500 conidia/mL).
C. Preparation of the closed system(s) and plant inoculation
C1. Closed systems in magenta boxes
Use magenta boxes or glass jars when a closed system that mimics growth in pots is desired (see Figure 1) (see General notes 4 and 5).
Figure 1. Lotus japonicus plants inoculated with (A) Fusarium oxysporum f. sp. lycopersici (Fol) and (B) F. solani strain K (FsK) and grown in glass jars purchased from glassware stores. In (A), the non-transparent lid of the jar is omitted, and a square piece of Parafilm is used instead to cover the jar mouth, allowing uniform light diffusion toward plants and gas exchange.
1. Place approximately 150–200 mL of substrate (sand/vermiculite mixture, 2:1) in magenta boxes/jars.
2. Autoclave containers filled with the substrate (121 °C for 60 min) prior to use to ensure aseptic conditions.
3. Perform plant transplantation under the laminar flow hood. Use Hoagland nutrient medium [12,13] to irrigate the substrate, prior to plant transplantation, when normal conditions for plant growth are desired. Use ~50 mL of nutrient medium per ~150–200 mL of substrate (see General note 6 and Troubleshooting 1). Transplant ~7–12-days-old L. japonicus plants into magenta boxes by using sterile forceps. Transplant 2–3 healthy plants per magenta box. Consider each magenta box as a biological replicate.
4. Close the containers with their lids and transfer them to the growth chamber until further handling (see General notes 7 and 8).
5. Perform fungal inoculation in the laminar flow hood to avoid contamination. Use fungal conidia as inoculum (as prepared in section B) in this closed system. Fungal conidia are easily quantified with the use of a Neubauer chamber (see General note 9 and Troubleshooting 2).
Note: For plant inoculations, a working concentration of 500 conidia/mL is used. Plants are inoculated by applying directly to the root 200 μL of inoculum (i.e., 100 conidia per root system).
6. Post fungal inoculation, close containers with their lids in the laminar flow hood and return them to the growth chamber until further experimental processing.
C2. Closed system in Petri dishes
Set the interaction system in square Petri dishes (see General note 10 and 11).
1. Perform plant transplantation under the laminar flow hood. Transfer ~7–12-day-old plants in square Petri dishes containing M medium (see General note 12) solidified with 0.4%–0.8% w/v agar under the laminar flow hood. Place Petri dishes containing the plants in the growth chamber at a ~45° angle (see General note 8).
2. Perform fungal inoculation in the laminar flow hood to avoid contamination from other microorganisms. In this closed system, use fungal hyphae from a previous fresh culture of the fungus in a PDA plate to inoculate the plants. Perform inoculations with a small fungal plug from a previous fresh fungal culture or a few fungal hyphae by scraping the surface of a previous fungal culture with a sharp sterile object such as the needle of a syringe. Inoculate at a spot close to the root, e.g., 1 cm underneath or close to the root tip (see Figure 2) (see General note 13).
Note: It is recommended to avoid inoculation with a large number of fungal propagules.
Figure 2. Square Petri dishes containing Lotus japonicus plants inoculated with Fusarium solani strain K (FsK). A few fungal hyphae from a previous routine fungal culture on solid medium was used to inoculate plants underneath the root tip using a sterile needle of a syringe. Scale bar: 1 cm.
3. Post fungal inoculation, close Petri dishes, seal them with Parafilm in the laminar flow hood, and return them to the growth chamber until further experimental processing. Place Petri dishes in a vertical position on the growth chamber shelves for optimal root growth.
C3. Closed system to perform temporal, in vivo observations
This section describes a non-destructive method for microscopical observations of colonized plants, appropriate for in vivo observations and cellular studies of the plant–fungal system using confocal microscopy (see Figure S1).
Set the plant–fungal interaction in square Petri dishes, underneath a thin (25 μm) gas-permeable membrane that allows prolonged microscopical observations (see General note 14). Perform all manipulations for system setup in the laminar flow hood to avoid contamination from other microorganisms.
1. Transfer pre-grown L. japonicus plants (~7–12 days old) to square Petri dishes containing M medium solidified with 0.4% w/v agar.
2. Remove a small portion of the solidified medium close to the root tip with a sterile needle from a syringe. In the space created, place a few hyphae from a previous fresh fungal culture grown on solid medium with a sterile needle from a syringe.
3. Fix the inoculum under 1–3 drops of a 0.4% w/v low melting temperature agarose solution.
Note: Allow the agarose solution to cool down before use.
4. Leave the agarose to solidify to ensure fixation of the inoculum (see Troubleshooting 3).
5. Pipette sterile water (~1 mL) around the roots and carefully cover the roots and the fungus with the gas-permeable plastic film, avoiding the entrapment of air bubbles underneath the film.
6. Seal Petri dishes with parafilm and wrap their lower part in aluminum foil to protect root tissues from light.
7. Place Petri dishes at a ~45° angle in a growth chamber (16/8 h light/dark photoperiod, 22 °C) until further handling.
8. Perform in vivo, temporal, and prolonged observations of the plant–fungal interaction directly on the Petri dish, using an upright (fluorescence) microscope by simply removing the Petri dish lid. No coverslip is needed for imaging. Tissues are directly observed underneath the gas-permeable plastic film [10,16] (see General notes 15 and 16).
D. Sampling plant tissues
1. Collect tissues for further processing, e.g., for staining and microscopical analysis. Collect plants at ~20 days post inoculation (dpi) to test for intraradical fungal colonization by different Fusaria strains. Explant plants and wash root tissues with water to remove substrate particles.
2. Stain tissues right after harvest or store them temporarily (at 4 °C, in water) for up to 2–3 days until staining or store them in ethanol solution for long-term storage (see General notes 17 and 18).
E. Staining of whole root/stem/leaf tissues to highlight fungal structures and mounting on microscope slides
This section describes how to examine Fusarium fungal colonization in L. japonicus root/stem/leaves. For this purpose, a similar staining protocol as that used for staining arbuscular mycorrhizal (AM) fungi colonized roots is described [17,18].
1. Prepare the stain by mixing the appropriate quantity of TB or CB with water, glycerol, and lactic acid (1:1:1 by volume).
Caution: Wear protective gloves/protective clothing/personal protective equipment during all steps of handling the above-mentioned dyes. Ensure adequate ventilation and avoid dust formation.
2. Clear tissues (i.e., remove cytoplasmic contents from roots) at the beginning of the staining process: cut roots into 1–5 cm segments and clear them with 10% w/v KOH for 10–30 min at 95 °C in a water bath (see General note 19).
3. Rinse roots 4× with deionized water, acidify roots with 1 N HCl for 20 min at RT, and subsequently stain the root segments either with 0.05% w/v TB or 0.05% w/v CB for 3–4 min at 95 °C and for extra ~20 min at RT (see General notes 20 and 21).
4. Wash tissues 4–5× with deionized water and place them at 4 °C in water overnight to remove excess dye.
5. Place tissues/tissue segments on a microscope slide using fine forceps, making sure that tissue segments do not overlap. Add a drop (or the necessary amount) of Polyvinyl-Lacto-Glycerol (PVLG) or glycerol (50% v/v) right before the left edge of the specimen. Place the left edge of the coverslip over the left edge of the sample and carefully lower the coverslip, covering the whole sample and avoiding air bubbles.
6. Observe prepared slides immediately or, preferably, place PVLG-mounted slides in an oven at 40 °C for 24–36 h or at 60 °C overnight to harden sufficiently and create a permanently mounted specimen [19,20] prior to microscopical observations (see Figure 3) (see General notes 22 and 23).
Figure 3. Lotus japonicus roots inoculated with different Fusaria strains and stained using three different stains: trypan blue (TB), chlorazol black (CB), and ink. The staining process for each stain is described in section E. (A–C) L. japonicus roots colonized by Fusarium solani strain K (FsK) at 27 dpi and stained with TB. The yellow triangles indicate FsK structures interacting with root hair or root epidermal cells. (D–F) L. japonicus roots colonized by Fusarium oxysporum f. sp. radicis-lycopersici (Forl) at 23 dpi and stained with CB. The yellow triangles indicate Forl spore-like structures on root epidermal cells. (G–I) L. japonicus roots colonized by Fusarium oxysporum f. sp. lycopersici (Fol) (G–H) or Forl (I) at 27 dpi and stained using the ink and vinegar method. The yellow triangles indicate (G) Fol propagules-like structures in a root hair cell, (H) Fol structures and propagules in a root cell, and (I) Forl hyphae on the epidermis of L. japonicus roots. Scale bars: 25 μm.
F. Embedding in agarose and sectioning
Prepare root and/or stem sections (transverse or longitudinal) to investigate the exact microbial location and progression within plant tissues and the changes occurring in either partner. Fungal location within delicate leaf tissues can be observed on intact tissues.
Prepare sections for microscopic analysis using a free-hand method. Pre-treat tissues for free-hand sectioning: soft and delicate tissues are easily destroyed if not embedded on a medium that acts as a scaffold, thus providing mechanical support and allowing maintenance of tissues/structures integrity. Such media can be agarose or paraffin (see General note 24).
This section describes how to perform sections by embedding tissues in agarose and performing sections by hand (see General note 25) (see Figure S2).
1. Prepare a 6%–8% w/v agarose solution. Melt the agarose solution and place it in a hot water bath until use.
2. Use commercial silicone rubber flat embedding molds (see General note 26). Segment roots/stems and place them in the mold cavities (one segment per cavity) with forceps.
Note: Align roots/stems in the center of each cavity (parallel to the length of the cavity), because after pouring the agarose, it is difficult to make further adjustments.
3. Allow the agarose solution to cool down near the point of solidification prior to embedding. Cover tissues with agarose by using a plastic Pasteur pipette cut at its end to ensure a larger opening, since the solution is viscous.
Note: Pay attention not to introduce air bubbles while pouring the agarose into the cavities containing the roots. Allow the agarose to solidify at room temperature.
4. After the agarose has solidified, remove agarose blocks from cavities and start sectioning the blocks containing the tissue segments using a sharp razor blade (see General notes 27 and 28).
5. Maintain sections until immediate use in deionized water (e.g., in a small Petri dish or on a microscope slide in 1–2 drops of water).
6. Place (stained) sections (see General notes 29 and 30) on a microscope slide with a drop of water or glycerol (50%–90% v/v) (see General note 31) and cover with a cover slip for observations under the microscope.
G. Embedding in paraffin wax and sectioning
This section describes how to perform sections by embedding tissues in paraffin wax and performing sections using a hand microtome (see General notes 32–34) (see Figure S3). To build a hand microtome, use a bolt and nut (~2 cm in depth).
1. Cut root/stem tissues in segments (making sure segments are a bit longer than the length of the nut) and maintain them in water until use.
2. Screw the bolt in the nut just for a few turns so that a space with closed bottom (a well) is created inside the nut.
3. Place some flakes of paraffin wax in 2 mL Eppendorf tubes and place tubes in a water bath at ~55 °C to melt the paraffin wax (see General note 35) (see Figure S3).
4. Position the hand microtome vertically (with the head of the bolt on the bottom) at a straight surface close to the water bath. Place a segment of root/stem tissue inside the nut and keep it with forceps (or with the aid of the forefinger) at its center and parallel to the longitudinal axis of the nut.
5. Quickly drop the melted wax inside the hand microtome. If necessary, place more melted wax to completely cover the well inside the nut. Allow the paraffin wax to harden at room temperature.
Note: Perform this step fast because melted wax hardens quickly at room temperature.
6. After the wax is solidified, use a sharp razor blade to remove and throw the excess wax from the upper top of the hand microtome (see Figure S3).
7. Turn the bolt slightly clockwise (less than a quarter of a full turn) to expose a small portion of the wax cylinder containing the root at its center. Perform a quick and precise section with a sharp razor blade by holding the blade perpendicularly to the wax cylinder containing the tissue (see Troubleshooting 4).
Note: Consider that the smaller the turn of the bolt, the thinner the section of the tissue. Perform a second section by slightly turning the bolt clockwise once again and so on.
8. Add 1–2 drops of water in a microscope slide and collect sections directly on the slide using fine point forceps or a pre-wet brush.
9. If necessary, remove the excess wax to expose the tissue section. This can be done with the aid of a dissecting (inoculation) needle under the stereoscope, as sections (especially root sections) are rather small and cannot be easily handled with the naked eye.
10. Stain tissues after embedding in paraffin and sectioning (see General note 30).
11. Place stained sections on a microscope slide with a drop of water or glycerol (50%–90% v/v) (see General note 31), and cover with a cover slip for observations under the microscope.
H. Microscopy
1. Observe prepared specimens under a compound light microscope and proceed with imaging using a microscope-mounted camera and the appropriate computer software for image acquisition (see Figure 3).
2. Use an epifluorescence microscope when fluorescent dyes are used to highlight specific plant tissues and compartments, fungal cells, and structures.
3. If accessible, use a confocal microscope to reject out-of-focus light from the image, for sharper images of cells and cellular structures without background fluorescence (see Figure 4) and for in vivo cellular studies.
Figure 4. Staining of endodermal suberization in Lotus japonicus root tissues inoculated with a Fusarium strain using the suberin-specific Fluorol Yellow (FY) stain. (A) FY-stained L. japonicus whole mount roots inoculated with the green fluorescent protein (GFP)-transformed isolate of Fusarium solani strain K (strain F9a) [10]. A single optical section is shown. (B) Maximum intensity z-axis projection of serial optical sections of the same root tissues shown in (A). L. japonicus plants (6 days old) were inoculated with F9a strain. Root tissues were harvested at 6 days post fungal inoculation and stained with FY using a lactic acid–based protocol, as described in Barberon et al. [21] and Shukla et al. [22]. Optical sections were collected using a Zeiss LSM800 system. Root tissues were imaged using the Ar laser band at 488 nm and a 500–550 nm emission window for FY fluorescence. Ep: epidermis; co: parenchymatic cortex; en: endodermis; white asterisks: passage cells. Scale bars: 50 μm.
I. Preparation of samples for fluorescence microscopy
Most cellular components are colorless; most molecules making up the cell do not absorb any specific wavelength of light, and therefore cannot be distinguished under the compound light microscope. A fluorescence microscope is an optical microscope that takes advantage of the physical properties of fluorophores (or fluorochromes): molecules that absorb light energy at a specific wavelength and re-emit it at higher wavelengths. This property of these molecules is called fluorescence, which can be visualized, recorded, and analyzed. Because the excitation and emission wavelengths differ, the absorbed and emitted by the fluorophore light are detectable in different areas (and therefore colors) of the spectrum and can thus be distinguished.
In a fluorescence microscope, the specimen is illuminated with a specific wavelength of light; the fluorophores within the specimen absorb the light and emit it at a longer wavelength, which typically falls within the visible or near-visible range of the spectrum. Specific filters separate the excitation light (high energy light) from the emission light (lower energy light) which is seen by the eye or captured by the detector. The detected fluorescence enables researchers to obtain spatial and functional information on cells/cellular components that have been stained/tagged with the fluorophore(s), among non-fluorescing material [23]. Some common types of fluorophores are:
• Many autofluorescent compounds, naturally occurring in plants, especially plant cell wall substances (examples of autofluorescent compounds in plants are phenols, lignin, suberin, etc.).
• Synthetic organic dyes (such as fluorescein)
• Biological fluorophores [fluorescent proteins such as green fluorescent protein (GFP)]
• Fluorescent dyes conjugated to target molecules (such as antibodies, proteins, peptides, etc.), which are used to label specific cellular components/compartments
• Quantum dots (nanocrystals that when excited, emit fluorescence at a wavelength based on the size of the particle) [23–25]
In this respect, in case experimental aims require the use of fluorophores to highlight specific plant tissues and compartments, fungal cells, and structures, a fluorescence microscope should be used (see Figure 4 and Troubleshooting 5).
In Table 1, a brief summary of common fluorophores used in the literature is presented, which may be used to highlight specific plant tissues and compartments, fungal cells, and structures.
Table 1. Commonly used fluorophores list. Brief summary of fluorophores commonly used in plant–microbe interactions (PMIs) to stain plant and fungal cells, compartments, and structures. *Solution preparations are indicatively given using information from only some of the citations corresponding to each fluorophore presented (**, the article from which information is drawn).
Fluorophore Use Solution preparation* Reference(s)
Acid fuchsin
To stain fungal structures of Serendipita indica (Piriformospora indica) in barley roots.
To stain fungal structures of S. indica (P. indica) in Arabidopsis thaliana roots.
0.01% w/v in lactic acid
[4]
[26] **
[27] **
Wheat germ agglutinin (WGA)-Alexa Fluor 488 conjugate
To stain chitin in the infection structures of Magnaporthe oryzae on rice leaf sheaths.
To stain fungal chitin and visualize S. indica (P. indica) structures in colonized barley roots.
10 μg/mL in 1× PBS (pH 7.4) containing 0.1% Triton X-100
[27]
[28] **
[29]
Calcofluor white
To visualize septa of S. indica (P. indica); specific dye for β-glucans and chitin.
In plants, it is able to bind cellulose in cell walls.
It exhibits selective binding to the cell walls of plants and fungi.
1.5 μg/mL
[30] **
[31]
Propidium iodide
It outlines cells in living plant tissue.
Used to assess root cell viability. It stains the nuclei of non-viable cells lacking intact plasma membrane (it penetrates the plasma membrane of non-intact plant cells, where it binds to DNA).
Used as a counterstain.
10 μg/mL in water
[10]
[28]
[32] **
SYTOX green, orange, and blue To assess plant cell viability in living plant tissues. The dyes are selective for non-viable cells. 250 nM in water [32] **
SynaptoRed C2 (FM4-64)
Endocytosis marker.
Membrane stain commonly used for dissecting vesicles trafficking in living plant cells.
To assess putative plant-derived membrane invagination events upon fungal accommodation.
Living cells internalize the dye into endomembrane structures.
3 μM in water
[10] **
[16]
[29] **
Fluorol Yellow 088
To stain suberin lamellae in plant root tissue.
To quantify suberized exodermal cells in cross sections.
0.01% w/v in lactic acid for sections; 0.05% w/v in methanol for whole roots
[33]
[34]
[35] **
Auramine O To stain suberin. 0.5% w/v [7] **
Berberine hemisulfate To stain Casparian bands. 0.1% w/v in lactic acid
[36]
[33] **
Nile red To stain suberin lamellae. 0.01% w/v in lactic acid (85%) saturated with chloral hydrate [37] **
Basic fuchsin To stain lignin. 0.2% w/v [7] **
Safranin O Used as counterstain to color lignified cell walls in botanical samples. To stain lignified tissues such as the xylem. To quantify lignin content. 0.2% w/v in 50% ethanol
[33]
[38]
[39] **
Acriflavine To stain lignin. Stains lignified cell walls green and unlignified cell walls red. 0.01% w/v aqueous solution [40] **
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
• Skiada et al. [10] Colonization of legumes by an endophytic Fusarium solani strain FsK reveals common features to symbionts or pathogens. Fungal Genetics and Biology 127: 60–74. (Figure 2, Figure 3, Figure 4, Supplementary Figure 3).
• Parts of the protocol have also been validated within the frames of the present manuscript. (Figures 1, 2, 3, 4, Supplementary Figures 1, 2, and 3).
General notes and troubleshooting
General notes
1. Alternatively, the plant material of choice may be used for examination (preferably a small-sized plant with a short life cycle). A filamentous fungus of choice may also be used as the microbial partner.
2. The duration of sulfuric acid treatment depends on seed age. Freshly harvested seeds require less scarification time.
3. In case a different fungal strain is used, modify accordingly the routine fungal culture and inoculum preparation.
4. In case magenta boxes are not an option, you may use common glass jars. These are relatively inexpensive and can be easily purchased in glassware stores.
5. Glass jars with a capacity of ~370 mL are a suitable choice for small plants such as L. japonicus. Keep plants in such containers for approximately one month. Afterward, stems reach the lid and space limitations occur.
6. Alternatively, M medium [14] or a nutrient medium of choice can be used.
7. In the case of jars, the lid may be loosely closed and further sealed with Parafilm to allow for gas exchange. If the light that reaches the plants is not sufficient due to the non-transparent lid, the lid may be omitted, and a square Parafilm piece may instead be used as cover (see Figure 1).
8. It is recommended to allow transplanted plants to rest for 2–3 days prior to fungal inoculation.
9. You may also use other types of inoculum such as fresh mycelium.
10. Chose Petri dishes as your closed system when, for example, you want (1) to examine the early stages of the plant–fungal interaction, (2) to visually record the exact time point of the interaction, (3) to visually inspect root system changes in relation to fungal inoculation, and/or (4) to perform in vivo microscopical observations in relation to time.
11. A drawback of the closed system in Petri dishes is that it does not allow prolonged maintenance of the plant–filamentous fungus interaction due to space limitations and overgrowth of the fungus to the detriment of the plant partner.
12. Alternatively, instead of M medium, a medium of choice may be used.
13. Inoculations with fungal propagules may be performed close to the plant at a spot of choice.
14. This system has been developed by Chabaud et al. [41] as a targeted arbuscular mycorrhizal (AM) fungi inoculation technique and has been adapted for confocal microscopy by Genre et al. [16]. It has been further modified by Skiada et al. [10] to meet the requirements of a targeted inoculation technique for plant–filamentous fungi interactions for temporal confocal microscopy observations.
15. A confocal microscope should be used for sharper images of cells and cellular structures.
16. Staining plant tissues for fluorescence microscopy may be performed directly on the Petri dish where the bipartite interaction is taking place. Excess stain is removed; no washing with water is performed afterward.
17. If you wish to stain tissues for microscopical imaging at a later time point due to time limitations, preserve tissues in 70% v/v ethanol solution, e.g., in labeled Falcon tubes at 4 °C. Rehydration of tissues is recommended prior to staining. Rehydrate tissues by simply immersing them in descending ethanol series for a few minutes and then in deionized water for a few extra minutes.
18. Tissues may be collected at the time point of choice, e.g., to test for intraradical fungal progression or for gene expression analyses. Tissues may also be collected and flash-frozen in liquid nitrogen for subsequent molecular analyses (e.g., for total RNA isolation and gene expression analysis). For such a type of analysis, store tissues at -80 °C until further processing.
19. Clearing time clearly depends on the tissue processed. Attention should be paid to leaf tissues, as these are delicate and can be easily destroyed in hot KOH. For L. japonicus root tissues, ~10 min in KOH at 95 °C is enough to clear tissues. Stem tissues require a longer clearing step; change KOH solution 2–3 times during this step.
20. Both TB and CB give good results in terms of intraradical staining of fungal cells. Chlamydospore-like structures of Fusarium, as described in Skiada et al. [10], are adequately stained. The ink and vinegar method as described in Vierheilig et al. [42] and Kosuta et al. [15] can also be used to stain Fusarium hyphae and structures in their interaction with L. japonicus root tissues. Spore-like structures of Fusaria are adequately highlighted with TB or CB. The thin Fusarium hyphae are better highlighted with the ink and vinegar method and are adequately observed under higher magnification (see Figure 3).
21. Santana et al. [43] proposed alternative, low-cost, and safer dyes as staining agents for the observation of AM fungi in root tissues. They proposed the use of organic food dyes, namely blue and red food coloring (5% v/v in commercial vinegar). Blue food dye gave better contrast between plant and fungal tissues and acted similarly in terms of color intensity and contrast to TB. Though these dyes work well with AM fungi, their efficacy for filamentous fungi staining is not investigated herein.
22. If time is limited, specimens may be observed immediately after preparation. It is recommended though to place PVLG-mounted specimens in an oven to harden completely before microscopical observations. This not only removes air bubbles from the specimen but also prevents residues from the mounting medium on microscope lenses.
23. PVLG mounting medium creates a permanent specimen, whereas glycerol mounting medium creates a semi-permanent specimen. A coverslip sealant is recommended to preserve slides over extended periods [44].
24. Free-hand sections, if prepared carefully, can be adequate not only for routine microscopical examination but also for microscopical imaging. It is advised to mount tissues before hand-sectioning in embedding media such as agarose and paraffin wax. Agarose is among the most common laboratory reagents, and paraffin wax can easily be purchased from local craft stores (e.g., in the form of flakes).
25. The process of embedding tissues in agarose and sectioning them is also described in Zelko et al. [36].
26. Small molds can also be created using aluminum foil as a cheap alternative to commercial flat embedding molds. Commercial flat embedding molds have the advantage of containing numbered cavities that are imprinted on agarose post-solidification.
27. If time is limited and sectioning of agarose blocks containing tissue segments is not feasible within the same day, blocks can be kept at 4 °C, in water, for a few days.
28. More sophisticated and expensive equipment such as a vibratome may be used to section agarose-embedded plant tissues, which allows for thinner and better-quality sections.
29. Tissues can be stained for fungal detection post embedding and sectioning since clearing and staining of root tissues result in fragile roots, which may collapse with further processing.
30. Tissues can be stained to label specific root elements such as the xylem, Casparian bands, etc., in case a more detailed analysis of the topology of fungal cells is required (see section I).
31. Adding more water to the glycerol solution mounting medium lowers the refractive index. In that way, you can adjust the refractive index of the mounting medium [44].
32. The method presented in Section G is commonly described in botany laboratory textbooks. A description of the procedure may also be found at:
https://gtac.edu.au/wp-content/uploads/2016/01/StainPlantStem_Sections_LabPreparation.pdf
33. This method is easier to use for L. japonicus stem sections, as root tissues are rather thin, though with proper and repeated practice, sections of root sections can also be achieved.
34. All materials for this technique can be purchased in local stores. Nuts and bolts can be purchased in local tool stores whereas paraffin wax can be purchased in local craft stores.
35. The melting temperature of paraffin wax is 54 °C, or it depends on the purchased product.
Troubleshooting
Problem 1: Increased humidity within the closed system used to grow the plant(s) and the fungal partner.
Possible cause: The substrate may be oversaturated if excess medium is added to the substrate. This will increase the humidity within the closed system, and plants will show overwatering signs.
Solution: We have observed that ~50 mL of volume of nutrient medium per 150–200 mL of volume of substrate ensures good humidity conditions within the closed system. Attention should be given to the nutrient(s) concentration and to the duration of plant growth within the closed system. The above depend on contextually experimental conditions and research questions.
Problem 2: Filamentous fungi used for plant inoculation may overgrow in closed systems (such as magenta boxes), and artificial adverse effects may be observed in plant(s).
Possible cause: Filamentous fungi tend to grow easily and fast in various substrates.
Solution: Avoiding inoculation with a high concentration of fungal propagules is recommended. For Fusarium, we propose inoculation with ~100 conidia per individual plant (per root system).
Problem 3: Fungal propagules dispersal may occur underneath the membrane, and the targeted inoculation is not ensured.
Possible cause: The water added underneath the membrane may entrain fungal propagules far from the inoculation point.
Solution: Avoiding inoculation with filamentous fungi by simply placing the inoculum in the vicinity of roots is recommended. Securing the inoculum with agarose prevents fungal propagules’ dispersal on the medium.
Problem 4: The first few sections performed with the hand microtome may not be of use.
Possible cause: The irregular wax shape and the position of the plant tissue at the edge of the bolt may contribute to a non-intact sectioning.
Solution: The following sections should be of use. The number of performed sections depends on the tissue length used and the “step size” of the turn of the bolt. The integrity of the sections is observed under the compound light microscope.
Problem 5: Some plant tissues may have strong background fluorescence, leading to difficulties during fluorescence imaging.
Possible cause: This may be attributed to certain autofluorescent compounds, naturally occurring in plant tissues.
Solution: Optimizing the excitation wavelength and emission window during fluorescence acquisition is recommended.
Supplementary information
The following supporting information can be downloaded here:
1. Figure S1. Setting up a closed system to perform temporal, in vivo observations of Lotus japonicus plants infected with a Fusarium strain.
2. Figure S2. Experimental procedure of embedding in agarose Lotus japonicus tissues infected with a Fusarium strain and sectioning the embedded tissues afterward.
3. Figure S3. Experimental procedure of embedding in paraffin wax Lotus japonicus tissues infected with a Fusarium strain and sectioning the embedded tissues afterward.
Acknowledgments
This work was supported by the Hellenic Foundation for Research and Innovation (H.F.R.I.) under the “3rd call for H.F.R.I. Research Projects to support Post-Doctoral Researchers” (Proposal Number: 07885) (to VS) and by the Research Committee of UTH (Programme No: 7322) (to KKP).
The authors would like to thank Constantinos Ehaliotis for the provision of facilities (Department of Natural Resources and Agricultural Engineering, Soil Science and Agricultural Chemistry Lab, Agricultural University of Athens, 75 Iera Odos, 11855, Athens, Greece), and Christina Nikolaou for aiding in picture taking of some of the images provided in supporting information.
This work has been partially validated in Skiada et al. [10] Colonization of legumes by an endophytic Fusarium solani strain FsK reveals common features to symbionts or pathogens. Fungal Genetics and Biology 127: 60–74.
Competing interests
Authors declare no competing interests.
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Publication history
Received: Sep 12, 2024
Accepted: Nov 21, 2024
Available online: Jan 5, 2025
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© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Plant Science > Plant cell biology > Tissue analysis
Plant Science > Plant immunity > Host-microbe interactions
Cell Biology > Tissue analysis > Tissue imaging
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5,187 | https://bio-protocol.org/en/bpdetail?id=5187&type=0 | # Bio-Protocol Content
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Locomotor Activity Monitoring in Mice to Study the Phase Shift of Circadian Rhythms Using ClockLab (Actimetrics)
AB Andrea Brenna
JR Jürgen A. Ripperger
UA Urs Albrecht
In Press, Available online: Jan 01, 2025
DOI: 10.21769/BioProtoc.5187 Views: 62
Reviewed by: Edgar Soria-GomezJesús Hernández FalcónMariana Astiz
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The authors used this protocol in eLIFE Jun 2024
Abstract
The circadian clock regulates biochemical and physiological processes to anticipate changes in light, temperature, and food availability over 24 h. Natural or artificial changes in white/blue lighting exposure (e.g., seasonal changes, jet lag, or shift work) can advance or delay the clock phase to synchronize physiology with the new environmental conditions. These changes can be monitored through behavioral experiments in circadian research based on the analysis of locomotor activity by measuring wheel-running revolutions. The protocol includes measuring the internal period length in constant darkness and administering nocturnal light pulses to mice kept either in light/dark conditions (LD 12:12, Aschoff-type II protocol) or continuous darkness (DD, Aschoff-type I). Here, we describe a step-by-step guide for researchers to analyze the mouse circadian clock using wheel-running experiments and ClockLab (Actimetrics) to quantify data.
Key features
• This protocol builds upon the method developed by Jud et al. [1], optimized for digital analysis using the ClockLab software.
• Step-by-step tutorial on measuring period length, analyzing periodograms, assessing general activity, and determining phase shifts (Aschoff Type I and II).
Keywords: Phase shift Aschoff-type II Aschoff-type I Phase delay Phase advance Circadian rhythms Nocturnal light
Background
Biological rhythms are adaptive physiological and metabolic processes, allowing virtually all organisms to anticipate changes in a light/dark cycle occurring over 24 h [2]. These processes are oscillating physiological responses to exogenous stimuli, namely zeitgebers or time-givers [3], such as light, temperature, and nutrients. These oscillations, driven by the above-mentioned stimuli, are defined as diurnal rhythms. The perception of white/blue light is one of the most important mechanisms for entraining the circadian clock to engage in diurnal behaviors [4]. Diurnal rhythms are virtually observable in almost all living organisms. In the absence of external stimuli (i.e., light), living organisms display a self-sustained biological rhythm called circadian (Latin: Circa Diem, around a day) with an internal period (tau, τ) slightly shorter than 24 h [5,6]. Since the divergence between circadian and diurnal rhythms can affect the survival rate, as observed in mice [7], living organisms must be entrained in the light/dark cycle (L:D) daily. Rodents are a well-known model for studying circadian rhythms by analyzing their rhythmic wheel-running activity [8]. Mice kept in diurnal conditions spend more time running on the wheel at night. Mice kept in constant darkness, by shielding them from external illuminating cues, display so-called free running or circadian rhythms, which are self-sustained in the absence of additional stimuli [1]. Nocturnal light can reset the endogenous circadian rhythms through a process called phase shift (ф) [9]. The direction of the phase shift depends on the clock’s temporal state. Light perceived in the early night promotes phase delays (namely, a delay in the activity onset). On the other hand, a light pulse late at night promotes phase advances (namely, anticipation in the activity onset), whereas light in the middle of the day does not alter the clock phase. The phase shift is manifested at the behavioral level with a change of locomotor activity onset (phase shift) the day after the light pulse. Here, we show a detailed protocol used for analyzing and quantifying the wheel-running activity of mice using ClockLab (Actimetrics) to determine period length, general activity profile, and phase shift of the circadian clock. We became aware that a step-by-step protocol that can guide researchers to perform such articulate experiments in a user-friendly way is not available online. Therefore, we propose a detailed version of the previously published one [1].
Materials and reagents
Biological materials
1. 3–6-month-old 129/C57BL6 mice
Equipment
1. Wheel-running facility
a. Soundproof ventilated chambers at constant temperature (22 ± 2 °C) and humidity (40%–50%)
2. Wheel-running cages
a. Cage (280 mm long × 105 mm wide × 125 mm high) (Tecniplast, catalog number: 1155M)
b. Stainless steel wire lid (Tecniplast, catalog number: 1264C116)
c. Stainless running wheel (diameter 115 mm) (Trixie GmbH, catalog number: 6083)
d. Magnet (Fehrenkemper Magnetsysteme, catalog number: 34.601300702)
e. Magnetic switch: Reed-Relais 60 (Conrad Electronic AG, catalog number: 503835-22)
3. Mouse housing
a. Water bottles (260 mL, 55 × 128 mm, polycarbonate, with silicone ring) (Tecniplast, catalog number: ACBTO262)
b. Bottle caps (Tecniplast, catalog number: ACCP2521)
c. Nestlets (5 × 5 cm) (EBECO)
d. Animal bedding (Schill AG, model: Bedding type 3–4)
e. Chow food (Kliba-Nafag, catalog number: 3432PX)
4. Illuminating system
a. Light bulb, 18 W (Mazdafluor, model: Symphony AZURA 965)
b. Light bulb mounting (230 V, 50 Hz, 0.37 A) (Mazda, model: Mx204-118)
c. Fan: accessories (CF-1212, 12 V=/500 mA) (Monacor, catalog number: 03.1670)
d. Luxmeter (Testo, GmbH & Co, 0–100.000 lux)
5. Computer hardware and software
a. Microsoft Windows PC (e.g., Dell, Intel Pentium III running Windows 2000 or higher)
b. Data acquisition board: National Instruments AMUX 64-T (fitted with 10-kΩ resistors)
c. RJ45 socket
d. PCI 6503 card National Instruments
e. National Instruments NI-DAQ software
f. ClockLab software package, Actimetrics
Note: ClockLab components can be purchased in a ready-to-use package from Actimetrics.
Procedure
Below, we describe the step-by-step procedure for characterizing the internal period length of mice and measuring the phase shift of their circadian clock by employing the wheel-running method. This method is largely used for studying modifications in the endogenous circadian clock of mice, reflected in an altered free-running behavior. In this section, we will describe two different approaches, named Aschoff-type II (phase shift measured in animals kept in diurnal conditions) and Aschoff-type I (phase shift measured in animals kept in constant darkness) [10]. We provide a step-by-step protocol that will facilitate users to analyze the period length (τ) and phase shift (ф) of the circadian clock. However, before starting to describe the procedure, the reader should become familiar with the following concepts that will be subsequently discussed:
• Actogram. Circadian rhythmicity in rodents can be measured by analyzing a general output like locomotor activity. In this specific approach, a sensor is connected to a wheel-running cage, which transmits the signal to the computer. The ClockLab (Actimetrics) software subsequently elaborates the data, which are displayed on the computer monitor (Figure 1A). Vertical bars appearing on the computer monitor are the readout of the number of wheel revolutions per time, and this specific graph is called an actogram. Each horizontal line represents one day. The height of each vertical black bar cluster indicates the sum of wheel revolutions happening in a defined amount of time (i.e., 10’). Mice are nocturnal animals, therefore, when they are kept under 12:12 h light/dark cycles, their activity is concentrated at nighttime (alpha), while the resting activity is in the daytime (rho) (Figure 1B). Under the 12:12 h light/dark cycle, resembling the solar day, alpha- and rho-phases are opposite in diurnal and nocturnal organisms [11]. Sometimes, scattered activity can be observed in the rho phase because the animal interrupts its sleep for a short time (Figure 1B, red box in the rho phase).
• Diurnal rhythm vs. free running. Diurnal rhythms are tied to the external stimuli (Zeitgeber). When the external stimulus is a light pulse (white or blue), we can refer to the time taken into consideration for analyzing the locomotor activity as Zeitgeber time (ZT). Within the 12:12 h light/dark cycle, ZT0 is defined as lights on, the beginning of the light phase, and ZT12 corresponds to lights off, the end of the light phase. Diurnal rhythms are equal to 24 h, and as mentioned above, the wheel-running activity is concentrated at nighttime. On the other hand, organisms kept in constant darkness can display the so-called free-running period or circadian rhythms. These rhythms may persist indefinitely in the absence of any synchronizing cue. The period length of these free-running rhythms is often not equal to 24 and differs from species to species. For instance, wild-type mice (C57BL/6Tyrc-Brd × 129S7) have an internal period length of 23.7 ± 0.1 h [12]. While in diurnal rhythms the temporal unit is the hour (ZT), in the free running we talk about the circadian hour (CT), which consists of quantifying the internal period length (tau) and dividing it by 24. During the first day of constant darkness, we can assume ZT = CT. Since circadian rhythms are manifested in constant darkness, we can talk about subjective day (CT0-12) and subjective night (CT12-24), where physiological processes reflect the diurnal profile. Figure 1C shows an example of the difference between diurnal and circadian rhythms.
• Entrainment. This is a process promoted by specific stimuli, such as light, food, and temperature, that induces the alignment between the endogenous clock and the external environment [13]. For instance, mice kept in constant darkness exhibit an endogenous period shorter than 24 h. However, when these rhythms are adjusted to a 12:12 h light/dark cycle, the endogenous clock aligns with the 24-h environmental rhythm after a few days. This process promotes the masking effect of diurnal rhythms over circadian ones. When constant blue/white light is applied to entrain the animals, we talk about the continuous mode. On the other hand, a short-term stimulus (minutes) given during the night can promote a phase shift of the circadian clock (see below). In this specific case, we talk about the phasic mode. The environmental signals that can entrain circadian clocks are called Zeitgebers [3] (Figure 1D, constant darkness vs. re-entrained light/dark cycle). When mice kept in constant darkness are re-entrained to a diurnal rhythm, we can observe a transient cycle [14]. It can be observed as a wheel-running revolution during the rho phase anticipating the activity onset, which can persist for a few days (Figure 1D, red box). Those transient cycles reflect the disequilibrium between the altered phase angle generated by the endogenous circadian rhythm of mice, which is < 24 h, and the entrained rhythm to the Zeitgeber (24 h) [11]. With the term “phase angle” (Ψ) we refer to the relationship between the timing of the biological clock and the timing of an external time cue [14].
• Phase shift. Phase shift (ф) refers to a shift in the animal’s activity onset as a consequence of a white/blue light pulse given during the night [15]. The phase response curve (PRC), whose shape varies depending on species and stimulus [14], displays the phase shift profile over 24 h when a light pulse is given every hour with mice kept in constant darkness. Figure 1E shows a typical light PRC of a nocturnal rodent. The phase curve can be divided into three parts: a phase delaying zone of the activity onset (CT12 to CT16), a phase advancing zone (CT18 to CT2), and a dead zone (CT2 to CT8, subjective day), observed when the stimulus does not affect the activity onset phase. Between CT16 and CT18 is the so-called singularity point where the PRC crosses the baseline (Figure 1E, blue star). If light hits exactly at that point (which is difficult to achieve), arrhythmicity is induced in the animal. The phase shift of the circadian clock can also be observed in diurnal conditions and the effect resembles what is observed in constant darkness (Figure 1E and F).
Note: Actograms are usually double plotted to show diurnal/circadian activity over multiple days, facilitating the identification of persisting rhythms (Figure 1F).
Figure 1. Digitalization and interpretation of actograms. A) Graphic representation of an actogram produced by ClockLab after elaborating on data obtained from the wheel-running cage and processed by the computer. The actogram is obtained from a published paper [16]. B) Representative single-plotted actogram displaying a mouse’s wheel-running activity showing activity bins (black vertical bars) concentrated at nighttime when mice are kept in 12:12 h light/dark conditions. The height of each vertical bar indicates the accumulated number of wheel revolutions for a given interval of time previously selected (e.g., 10 min). Each horizontal line corresponds to one day. The rho- and alpha-phases indicated on the top of the actogram refer to rest and activity, respectively. The red box in the light phase indicates the scattered activity of a mouse during the resting phase in the daytime. The red box in the dark phase indicates that the mouse is resting at nighttime. C) Representative single-plotted actogram shows the different distribution of mouse activity when it is kept in diurnal (12:12 h light/dark) or circadian (free running) conditions. The Tau (τ) in a 12:12 h light/dark condition is approximately 24 h, while the one displayed by mice kept in constant darkness is slightly shorter. Since the computer timer is set on the 24-h day cycle, the wheel-running activity in constant darkness looks like a slope. D) The single-plotted actogram displays in the red box the mouse activity in the daytime as a consequence of the re-entrainment from the constant darkness to the 12:12 h light/dark cycle. This event produces a transient cycle. E) The phase response curve (PRC) for nocturnal rodents, produced in circadian conditions (constant darkness) [1]. The gray and black bars below the PRC indicate subjective day and night, respectively. The X-axis shows the circadian time (CT) at which the researcher applied the light. The Y-axis displays the amplitude in hours of the phase shift (ϕ). Light pulses administered between CT11 and CT16 provoke a phase delay (negative values), whereas light pulses between CT19 and CT3 generate phase advances (positive values). A light pulse given between CT4 and CT10 does not produce any phase shift (dead zone). We can observe the inversion point in the phase shift response starting at CT16 and reaching the singularity (blue star) at CT18. F) The double-plotted actogram shows examples of a phase delay (middle actogram), phase advance (right actogram), and no delay (left actograms) following the Aschoff-type II (upper actograms, diurnal conditions) and Aschoff-type I protocol (lower actogram, circadian conditions). The yellow stars indicate when the light was applied. For more details, please check the original paper [16].
A. Mice preparation and experimental setup
1. Plastic cages with steel running wheels must be prepared with bedding and nesting material (General note 1) (Figure 2A).
2. Weigh mice between 3 and 5 months old and check to evaluate their health (General note 2). Weight and health conditions should be noted on an appropriate scoresheet approved by the veterinary office (Document S1).
3. House mice individually in wheel-running cages with access to food and water ad libitum (Figure 2B).
4. Wheel-running cages are provided with a small magnet nestled in a plastic disc that rotates when the wheel moves (Figure 2C). The signal is transmitted to the computer through a connector plugged close to the plastic disc, and the rotations are visualized by the clock lab software (Figure 2D). Plug the connector and test the magnet before starting the experiment (General note 3). Each cabinet can contain a maximum of 12 boxes. Cabinets are soundproof and ventilated (General note 4).
5. Set the timer to have light/dark (LD) cycles of 12:12 h. The timer is mounted outside the cabinet (Figure 2E). Lock the cabinets (Figure 2E) and start the recording. The illumination is ensured by two light bulbs (1,000 lux, Figure 2F) mounted on the ceiling of the cabinet (General notes 5 and 6). The light intensity is confirmed by a luxmeter (Figure 2G).
6. During the first day of the current experiment, conditions (L:D, D:D), number of recorded days, mouse genotype, age, and sex need to be annotated before the experiment into an appropriate sheet (Document S2).
7. The experimenter should be particularly careful in monitoring mice activity during the first days (General note 7).
B. Wheel running activity recording and validation of the free-running period
1. Mice should be entrained to the L:D 12:12 cycles for at least 10 days to adapt to the isolation cabinet and the diurnal cycle. At this stage, the entire activity will be confined to nighttime since mice are nocturnal animals, and the actogram will show two separate phases: a resting (rho) and an active phase (alpha) (Figure 1B).
2. After at least 10 days, mice can be released in constant darkness, and a free period will appear on the actogram like a slope (Figure 1C). If the experimenter wants to measure only the free period, they can turn off the illuminating system anytime, after ZT12. Turning the light off after ZT12 would allow mice to enter the free-running period the day after. The illuminating system should be turned off at ZT10 to validate the Aschoff-type II protocol.
3. The free-running period should last at least 10 days. Once a stable free-running rhythm is established, the experimenter can determine the internal period length and the overall activity as revolutions/day. The free-running period of wild-type mice should be less than 24 h. Since the computer clock is set to 24 h, the free running will appear on the actogram like a slope (Figure 2D).
4. Mice are re-entrained to the light/dark cycle for at least 10 days (i.e., Figure 1D) before being ready for the phase shift of the circadian clock protocol.
Figure 2. Essential tools for the experimental procedure. A) Representative cage with nesting and bedding materials. B) A photo of the cage showing the food and water configuration. C) Example of a wheel-running cage connected via a magnetic switch to the system recording wheel revolutions. On each rotation of the running wheel, the magnet, embedded in the plastic disc, activates the magnetic switch, opening and closing. The magnetic switch cable sends the info to the computer, which elaborates the data displaying the number of wheel-running revolutions in the actogram. D) Representative actogram displayed on the computer when the info obtained from the magnetic switch is elaborated by ClockLab. E) The external timer allows the user to set the appropriate light/dark cycle, avoiding multiple cabinet openings, which might eventually drop the internal temperature. The red arrows point to the timers. The blue arrows point to the power generator. F) Example of a cabinet interior and illuminating system. G) Example of a luxmeter that can be used for measuring light intensity.
C. Phase shift of the circadian clock
The phase shift of the circadian clock can be measured with two different protocols: Aschoff-type II and Aschoff-type I. Applying the type II protocol, we give light pulse at ZT14 and ZT22 followed by releasing mice in constant darkness and measure the angle between the free-running and diurnal periods. Applying the type I protocol, mice are already released in constant darkness. Therefore, we measure the endogenous circadian hour and give the light pulse at CTs 14, 22, and 10. A schematic example of the protocol is shown in Figure 3. For more details about how to calculate the proper CT, please see the data analysis section.
Figure 3. Schematic representation of the Aschoff-type II and Aschoff-type I protocol for measuring the phase shift of the circadian clock. After ten days under diurnal conditions (blue line; L:D 12:12), mice receive light exposure (Aschoff-type II) at ZT14 (2 h after lights off) and ZT22 (2 h before lights on). The control at ZT10 can be achieved simply by switching off the light at that specific time point. Following the light pulse, mice are kept in constant darkness for ten days (free running, black line). After ten days under circadian conditions (grey line), the appropriate CT14, CT22, and CT10 are determined, and light is applied (Aschoff-type I). Mice are then monitored for another ten days in constant darkness (black line).
Why use two different protocols? The Aschoff-type I can be applied to animals with a stable free-running rhythm when kept in constant darkness. We can apply only the Aschoff-type II protocol when animals show arrhythmic free running (i.e., Per1ko/Per2ko mice). Due to the arrhythmicity, it is impossible to calculate the CT based on the endogenous circadian hour. Additionally, for fast screening of mutant animals that might display aberrant phase shift responses to light pulses, the Aschoff-type II protocol is easier to apply because we can give the light pulse to all animals simultaneously. However, if mutant mice show a regular free-running period, the Aschoff-type I protocol needed to be applied as well. The advantage of the type I protocol is that the phase shifts can be easily determined as the difference between two almost parallel lines. However, for every individual animal, the time of light application has to be calculated based on its period (see below).
Aschoff-type II
1. Mice should be entrained into the L:D 12:12 cycles for at least 10 days to adapt to the isolation cabinet and the diurnal cycle. After 10 days, 15’ of a light pulse can be applied at ZT14 (phase delay), and mice can be released for another 10 days in constant darkness to show a stable free-running period.
2. Mice are re-entrained into the light/dark cycle for at least 10 days before being ready for the phase shift of the circadian clock protocol.
3. After the re-entrainment, 15’ of a light pulse can be applied at ZT22 (phase advance), and mice can be released for another 10 days in constant darkness to show a stable free-running period.
4. Mice are re-entrained into the light/dark cycle for at least 10 days before being ready for the phase shift of the circadian clock protocol.
5. After the re-entrainment, 15’ mice can be released for another 10 days in constant darkness directly at ZT 10 (no phase delay).
Aschoff-type I
1. The circadian hour will be calculated based on the circadian free-running period (see the next section) obtained from the previous protocol (Aschoff-type II, step 5). Then, the specific time point will be calculated based on the CT.
2. 15’ of a light pulse can be applied at CT14 (phase delay), and mice can be kept for another 10 days in constant darkness.
3. An additional 10 days of constant darkness ensures that the effect of the previous light pulse is eliminated.
4. 15’ of a light pulse can be applied at CT22 (phase advance), and mice can be kept for another 10 days in constant darkness.
5. An additional 10 days of constant darkness ensures that the effect of the previous light pulse is eliminated.
6. 15’ of a light pulse can be applied at CT10 (no phase delay), and mice can be kept for another 10 days in constant darkness.
7. Mice can be re-entered in a light/dark cycle if they are needed for other experiments.
Data analysis
A. How to open an actogram file in Clock Lab
1. Open Clock Lab.
2. Go to Open, search for the actogram file that can be read by ClockLab, and click on it (Figure 4A).
3. The actogram appears in the Clock Lab (Figure 4B).
4. The user can choose between a single (Figure 4C) and a double (Figure 4D) plot actogram. To double-plot the actograms, the user can go to the tool settings and select double-plotting.
B. Analysis controls
Start/end dates: The menu contains a list of the dates (month-day-year) for which data were collected, together with the number of each day that appears on the actogram on the Y-axis (Figure 4E). These menus can be used to set the range of data from which all calculations and graphs are made.
Start/end hour: They determine the time frame window (hours) within which the actograms are analyzed. This control is particularly important for the actogram display. The user can set the hour (i.e., hour 1: 7:00; hour 2: 24:00) to match the onset and offset (i.e., 7:00 light on, 19:00 light off) to have the rho and alpha phases properly distributed (Figure 4F).
Tau: It influences the appearance of the actogram and the activity profile. The default Tau is exactly 24 h (Figure 4G). If the Tau is set with the time corresponding to the mouse free-running period, the actogram will not appear like a slope anymore, but it will look straight and L:D will look like an advanced slope (Figure 4H).
Bins: The user can set the bin width, namely the window of time taken into account for measuring the wheel-running revolutions (Figure 4I).
Type: Four types of actograms can be displayed (percentile distribution, even distribution, threshold, scaled). The choice of the type will influence the appearance and size of the black bars associated with the count of wheel-running activity (Figure 4J). For further details, please visit the appropriate website (https://actimetrics.com/products/clocklab/).
Onset/offset: The red dots show the results of the automated wheel-running activity onset. They are necessary indicators for drawing the regression line used for measuring the internal period length. Given the onset and offset time, the software can calculate at a rate of 95% correctly for most data. If the red dot is misplaced, it can be corrected simply by shift-clicking on the appropriate point within the line for that day (Figure 4K).
Figure 4. Step-by-step guide for measuring chronobiological parameters using ClockLab. A, B) Representative screenshots showing how to load an actogram on ClockLab. C) Difference between single and D) double-plotted actogram. E) Representative screenshot showing how to select the days for analyzing data. F) Representative screenshot showing how to center the actogram to have half of it showing the light phase and half of it showing the dark phase. In this case, the light was on at 07:00 and off at 19:00; therefore, setting 07:00 in the hour box would let the actogram start when the light is on, and the onset activity would appear in the middle of it. G) Representative screenshot showing how to select the tau. Tau:24 means that the mice profile defined by their wheel-running activity is distributed over 24 h. H) Example showing how manipulating the tau can change the shape of the actogram. If the tau is set to 24 h, the actogram looks straight in diurnal conditions and as a slope with anticipating onset in circadian conditions. If we set the tau at the circadian period in constant darkness, then the actogram in diurnal conditions will look like a slope delaying the onset every day, while the actogram in constant darkness will look straight. I) Representative screenshot showing how to select the window of time displaying the number of wheel-running revolutions. Bin:1 means how many revolutions per minute. J) Representative screenshot showing how to modify the appearance of the actogram by selecting the more appropriate “type.” K) Representative screenshot showing how to select the onset and offset time. The red dots (their size was increased to make them visible) displayed on the actogram onset of the mouse activity profile for each day are adjusted accordingly.
C. Features
1. Period length
Mice must be kept for at least 10 days in constant darkness to quantify the free-running period length (τ). Then, the red dots displaying the activity onset need to be properly assigned at the beginning of the daily wheel-running activity in constant darkness. Subsequently, a right-click on the mouse will show a drop-down menu (Figure 5A). The user should click fit1. The tool generates a least-squares fit to a group of the points (each point is one day) displayed in the actogram window. Finally, it draws a line of the selected following days of activity, excluding the first two (transient cycles). The endogenous period length (τ) will be determined from the regression line drawn through the activity onsets [17] (Figure 5B). Along with the fitted line and tau, we can analyze (Figure 5B, red box):
• Error: The standard deviation of the horizontal distance between the onset activity points and the regression line.
• Mean: The mean time in hours calculated for the regression line, considering the number of days taken into account for calculating the tau.
The calculated period length can be used to compare different genotypes or pharmacological conditions.
2. Periodogram
A periodogram, namely the periodogram of Enright [18], consists of an analysis employed for short-term rhythms. It represents one of the options offered by ClockLab for measuring accurately the internal period length. The periodogram is calculated within a period set by the tau (i.e., 24 h) displayed on the x-axis of the graph. The data are cut into segmental periods (i.e., data analyzed within a window of two consecutive hours) within 24 h. The same segments, also called modules, from several days of analysis (at least seven consecutive days) are averaged together to give an activity profile, and the standard deviation of the profile is calculated. Following the Enright model, circadian data are plotted to provide a quotient of variance across the different modulo segments. Each modulo corresponds to a specific circadian period. The modulo with the largest value determines the dominant period length. The user should select from the “start-end” boxes the range of days that have to be analyzed. Then, on the main window, select analysis and finally periodogram (Figure 5C). The blue trace will show peaks of activity calculated by averaging several days within each segment of two hours composing the entire tau (24 h). The highest peak will determine the period length. Of note, we reported two cases—one where the periodogram measured the period in constant darkness (23.55 h) and the other where the period was measured in L:D conditions (24.05) (Figure 5D). The mouse analyzed was the same. Together, these results indicate that the periodogram is an efficient tool for precisely measuring the internal period. The importance of this model is given by the fact that it can also display ultradian rhythms [19] compared to the least-squares regression fit (Figure 5B), recurrent periods, or cycles repeated throughout a 24-h day. Figure 5D indicates with red arrows a recurring ultradian rhythm with a period of circa 12 h (masked by the circadian rhythm at 24 h). There are three different statistical analyses based on different formulas or algorithms used for implementing the calculation of the internal period length within a periodogram. These can be selected by clicking on Type (Figure 5E, red arrow):
• Chi-squared (χ2) periodogram: It is calculated after the method of Sokolove and Bushell [20]. χ2 is defined as a ratio of the variance observed at a specific period segment compared to the total average variance in the data set. χ2 values increase with the amplitude of the rhythm. If the amplitude is generally low, the χ2 becomes a less sensitive tool.
• Fourier-periodogram (F): It is calculated after the method of Dörrscheidt and Beck [21]. F analysis implies that any modulo can be approximated by a series of sine and cosine waves of differing period, amplitude, and phase. Therefore, the modulo is transformed into a series of simple waveforms, with coefficients determined based on the goodness of fit to the data.
• Lomb-Scargle periodogram: It is calculated after the method of Lomb [22]. It is an adaptation of the F-periodogram for datasets with missing values or irregular data [23].
The user can select the statistical analysis (chi-square, F, Lomb-scale) on the box type. In our example, the selected chi-square will appear as a green line (Figure 5D), and the period is measured considering the spectrum that is crossing the Chi-squared line.
3. Activity profile
The activity profile displays the distribution of the activity onsets (wheel-running revolutions counts) over 24 h, averaging the range of days taken into account. The user can obtain an activity profile of the selected range of days, by clicking on Analysis and then Activity profile. The software will calculate the distribution of the mouse's general activity divided over 24 h. As shown in Figure 5F, in diurnal conditions, the mouse is active starting from ZT12 to ZT0 (alpha phase) and resting from ZT0 to ZT 12 (rho phase). Interestingly, as mentioned before, during the resting phase, mice can show scattered activity, which is also displayed by the activity profile. On the other hand, between ZT18 and ZT22, mice show a “siesta” moment, where their activity drops before starting again until the end of the dark phase. The shading around the activity profile trace indicates the standard deviation for each point.
Users can find in the activity profile menu the following items (Figure 5F, red box):
• Fit: Select the phase angle to have the activity profile phased to the 24 h (solar day).
• Units: The user can set the x-axis in degrees (t/Tau*360), hours, circadian hours, %, or Tau.
• Sin fit: The red line is a sine fit to the waveform. Parameters describing the fit are shown in the table to the left, including amplitude, phase, and mean.
• Amplitude: Delta between MESOR (mean of the circadian rhythm) and the peak of the sinusoidal-shaped circadian rhythm (Figure 5G).
• Period (24 h): The period is used for measuring the phase angle (it is usually 24 h). The period is the distance between two peaks of the sinusoidal-shaped circadian rhythm (Figure 5G).
• MESOR: Midline statistic of rhythm; it is a rhythm-adjusted mean to the individual rhythm (Figure 5G).
• F statistic: It represents a measure of the robustness of circadian activity rhythm. Increasing values indicate a stronger rhythm, whereas decreasing values indicate a weaker rhythm.
• Estimate of p-value: For such a robust rhythm as shown here, it is arbitrarily close to zero.
• Mean: Places a horizontal line in the activity profile at the mean or median of the graph. This control also determines whether a mean or median line appears in a batch printing of activity profiles.
• Export: Export is part of the activity profile analysis tool. It displays counts for each day in the MATLAB command window and saves the counts to a spreadsheet-compatible file like Excel. The file also contains the raw data (time and counts/day) of the activity profile plot.
Figure 5. Step-by-step guide for measuring the circadian period length using ClockLab. A, B) Screenshots showing how to measure the period length based on the activity onset calculated on the assigned red dots (their size was increased to make them visible) of eight following days. C) Screenshot showing a representative periodogram selected from the analysis tool. D) Example of two different periodograms. The upper one displays the period length of a mouse kept under diurnal conditions (12:12 h light/dark), and the lower one displays the period length of the same mouse kept under circadian conditions (constant darkness). The red arrows indicate ultradian phenotypes that appear with a periodicity of 12 h (12, 24, 36 h). Note that at 24 h, the phenotype is masked by the dominant circadian period. The green line indicates Chi-squared. E) Screenshot showing how to select the appropriate statistical test for the periodogram. In the figure, the Chi-squared periodogram is shown. F) Screenshot showing a representative activity profile graph of a diurnal mouse. An activity profile is an option contained in the analysis tool. In the red box, all parameters analyzed are displayed. G) A schematic representation that shows the graphical meaning of some relevant parameters like the period, MESOR, and amplitude. For the explanation, read the appropriate section in the text.
4. Phase shift
a. Aschoff-type II
The actogram should display at least 10 consecutive days in diurnal and 10 consecutive days in circadian conditions. When the onset activity time is set for mice in both L:D and constant darkness conditions, the least-squares regression fits the onsets that have been calculated for the diurnal and circadian rhythms. Usually, with fit1, we draw a regression line for the following days in constant darkness, excluding the first two days to avoid the effect caused by the transition cycles. The fit2 regression line is dragged to the seven following days of the previous diurnal cycle (Figure 6A). When the user measures the phase shift, they need to be sure to point the mouse cursor at the part of the angle corresponding to the second day after the light pulse (yellow star, Figure 6B) to have a better-optimized phase shift. Then the user can move the cursor to the angle formed by the two regression lines and click on phase shift (Figure 6C).
b. Calculation CT
To perform the Aschoff-type I protocol, it is necessary to calculate the subjective phases (CT, or circadian time) of an animal’s rhythm based on the individual organism’s free-running rhythm. To that extent, the first thing to do is to calculate the circadian hour by dividing the internal period length τ (see section C1 of Data analysis) by 24 the day before the phase shift experiment (Day A):
1 circadian hour = τ/24
For instance, if the free-running period is 23.7 h, calculated in Day A, the circadian hour will be 0.987 (23.7/24). It has to be taken into account that for each day spent in constant darkness, CT12 diverges further from ZT12. On the first day in constant darkness, we can assume they are almost the same, but each day, the divergence between the internal and external tau increases (i.e., 24 - 23.7 = 0.3 h of divergence for each day). Thus, to be precise about calculating the proper CT10, CT14, and CT22, we can proceed as follows.
• The researcher has to calculate the CT12 of the next day (Day B) by applying the following formula:
CT12 Day B = CT12 Day A + τ - 24 h
For instance: CT12 Day B = (0.987*12) + 23.7 - 24 h → CT12 = 11.544 h
• To define a circadian time from CT0 to CT12, one can apply the following formula:
CTX0-12 = CT12Day B - (X*1 circadian hour) → [X = CT12 - CTx]
For instance: CT10 = 11.544 - [(12 - 10)*0.987] → CT10 = 9.57 h
• To define a circadian time from CT12 to CT24, one can apply the following formula:
CTX12-24 = CT12Day B + (X*1 circadian hour) → [X = CTx - CT12]
For instance: CT14 = 11.544 + [(14 - 12)*0.987] → CT14 = 13.518 h
Aschoff-type I
The actogram should display at least 10 consecutive days in constant darkness before and after the light pulse. When the onset activity time is set for mice before and after the light pulse, the least-squares regression fits the onsets that have been calculated (we proceed as we did for the Aschoff-type II protocol). We double-click on the second day after the light pulse, and we calculate the phase shift (yellow star, Figure 6D). It is important to note that it is easier to visualize the phase shift of the circadian clock applying the Aschoff-type I protocol since when the phase shift happens, fit1 and fit2 run parallel (Figure 6D).
Figure 6. Step-by-step guide for measuring the phase shift of the circadian clock using ClockLab. A–C) Representative screenshots showing how to calculate the phase shift of the circadian clock when the Aschoff-type II protocol is applied. D) Representative screenshots showing how to calculate the phase shift of the circadian clock when the Aschoff-type I protocol is applied. The yellow stars indicate where to point the cursor before clicking phase shift.
Statistical analysis
Statistical analysis can be performed using GraphPad Prism6 software. Depending on the data type, either an unpaired t-test or one- or two-way ANOVA with Bonferroni or Tukey’s post-hoc test can be performed. Values considered significantly different are highlighted [p < 0.05 (*), p < 0.01 (**), or p < 0.001 (***)].
Data compared via one-way ANOVA with post hoc Bonferroni’s corrections requires repeated-measures multiple comparisons among columns. Data (normal) distribution must be assessed, and accordingly, parametric or non-parametric tests must be applied. All data must display the mean ± standard error of the mean (SEM) and individual values. The suggested number of mice should be at least six per experimental group.
Validation of protocol
This protocol or parts of it have been used and validated in the following research articles (among others):
• Brenna et al. [24]. Cyclin-dependent kinase 5 (CDK5) regulates the circadian clock. eLife. (Figure 2)
• Brenna et al. [16]. Cyclin-dependent kinase 5 (Cdk5) activity is modulated by light and gates rapid phase shifts of the circadian clock. eLife. (Figure 1)
• Chavan et al. [25]. Liver-derived ketone bodies are necessary for food anticipation. Nat Commun. (Suppl. Figure 2)
• Schmutz et al. [26]. A specific role for the REV-ERBα-controlled L-Type Voltage-Gated Calcium Channel CaV1.2 in resetting the circadian clock late at night. J Biol Rhythms. (Figures 1 and 2)
Advantages and disadvantages
To date, the wheel-running activity is still the best method for extrapolating important information about diurnal/circadian behaviors such as free-running period, phase angle of entrainment to the LD cycle, period amplitude, and daytime running activity. Although numerous methods exist to measure the phase shift of the circadian clock (i.e., monitoring general activity using photo beams or the core body temperature by implanting sensors [27]), the wheel-running activity is still the less invasive approach. Compared to the general activity measurement via photo beams or core temperature, wheel-running measures only voluntary activity and not other activities such as grooming. Due to this, the wheel-running shows clearer onsets and offsets of activity, making the interpretation of an actogram easier. However, there are several points to take into consideration. The lighting conditions should be adjusted via a timer without opening the box. The isolation cabinets are well-ventilated to avoid overheating. However, light is a stronger and more immediate Zeitgeber than temperature. Therefore, the observed temperature variations under LD conditions can be neglected. Under constant darkness conditions, the temperature excursion during the day should be minimal since environmental temperature cycles can sustain peripheral circadian clocks [28]. Mice are kept in single caging for a certain amount of time. Although individual cages are close to others, and mice can still smell each other, long exposition to solitude might eventually affect their mood (i.e., stereotypical behaviors). ClockLab provides a wide range of options for measuring all the circadian parameters described above. However, any option comes with strengths and weaknesses. Please read the review [29], which gives more insights about the topic.
General notes and troubleshooting
General notes
1. The experimenter should not use excessive bedding to avoid wheel blockage while the mouse is on it.
2. It is important that age, gender, and weight match as much as possible to avoid variability that might affect the mouse’s wheel-running activity [30–32].
3. Once the connector is plugged into the cage, manually rotate the wheel and check on the computer if the ClockLab software is counting the revolutions.
4. Please turn off the room's light and check whether the cabinet is properly isolated (no light from the inside should be visible).
5. The light intensity is an essential factor. Therefore, confirm the proper light intensity using a luxmeter. Regular gray/black mice can perceive light up to 1,000 lux. Albino mice should be treated more carefully since they are more sensitive to light. The color temperature of the bulb is an important factor, measuring the visual “whiteness” of the light, and its unit is degrees Kelvin (K). Blue light, which is cold, on the other hand, displays a high color temperature and ranges. The necessary signal for entraining mice to the light/dark cycle is contained in the white light with natural daylight at temperatures between 6,000 and 7,000 K. The bulb described above has a luminous flux of 1,000 lumens and a color temperature of 6,500 K.
6. The time set on the timer should correspond to the actual zone time corresponding to the experimenter's location.
7. During the first three days of the experiment, the experimenter should monitor the actogram produced by the ClockLab software to ensure that all the connectors are recording and the proper light and dark conditions are working fine.
Troubleshooting
Problem 1: Light dispersion from the cabinet.
Possible cause: Door seals damaged.
Solution: Replace door seals.
Problem 2: The computer is not detecting wheel-running revolutions.
Possible cause: The magnet or magnetic switch is damaged.
Solution: Replace the magnet or magnetic switch.
Problem 3: The computer displays few or no wheel-running activity for a specific mouse on the actogram.
Possible cause: The wheel is stuck with bedding, or the mouse is sick.
Solution: Open the cage during the light phase and check whether the problem is the wheel or the mouse.
Problem 4: The computer displays activity bouts anticipating or delaying the external onset/offset (light on/light off).
Possible cause: The timer is not working properly
Solution: Check at the time of day when the light should be turned on and off to confirm that the timer is working. If it is not working, replace it.
Problem 5: The phase shift of the circadian clock (Aschoff-type I) did not work as expected in wild-type mice.
Possible cause: The CT calculation was wrong.
Solution: Check all the parameters again and perform new calculations.
Supplementary information
The following supporting information can be downloaded here:
1. Document S1. Score sheet
2. Document S2. Wheel running datasheet
Acknowledgments
This work was supported by the Swiss National Science Foundation (SNF) 310030_219880/1 to UA. This protocol is used in the following paper: Brenna et al. [16]. Cyclin-dependent kinase 5 (Cdk5) activity is modulated by light and gates rapid phase shifts of the circadian clock. eLife. e97029.
Competing interests
No competing interests were declared.
Ethical considerations
The cantonal veterinarian's office approved all protocols of the Fribourg state (license numbers: 2021-19-FR).
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Article Information
Publication history
Received: Sep 20, 2024
Accepted: Nov 28, 2024
Available online: Jan 1, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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5,188 | https://bio-protocol.org/en/bpdetail?id=5188&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
A Bio-protocol resource
This is an In Press version of the protocol that has not yet been assigned to an issue.
Peer-reviewed
Protocol to Mine Unknown Flanking DNA Using PER-PCR for Genome Walking
ZY Zhou Yu *
DW Dongying Wang *
ZL Zhiyu Lin *
HL Haixing Li
(*contributed equally to this work)
In Press, Available online: Dec 26, 2024
DOI: 10.21769/BioProtoc.5188 Views: 29
Reviewed by: Alba BlesaMichael Kotik Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Molecular Genetics and Genomics Mar 2024
Abstract
Genome walking, a molecular technique for mining unknown flanking DNAs, has a wide range of uses in life sciences and related areas. Herein, a simple but reliable genome walking protocol named primer extension refractory PCR (PER-PCR) is detailed. This PER-PCR-based protocol uses a set of three walking primers (WPs): primary WP (PWP), secondary WP (SWP), and tertiary WP (TWP). The 15 nt middle region of PWP overlaps the 3' region of SWP/TWP. The 5' regions of the three WPs are completely different from each other. In the low annealing temperature cycle of secondary or tertiary PER-PCR, the short overlap mediates the annealing of the WP to the previous WP site, thus producing a series of single-stranded DNAs (ssDNA). However, the 5' mismatch between the two WPs prevents the template DNA from synthesizing the WP complement at its 3' end. In the next high annealing temperature cycles, the target ssDNA is exponentially amplified because it is defined by both the WP and sequence-specific primer, while non-target ssDNA cannot be amplified as it lacks a binding site for at least one of the primers. Finally, the target DNA becomes the main PER-PCR product. This protocol has been validated by walking two selected genes.
Key features
• The current protocol builds upon the technique developed by Li et al. [1], which is universal to any species.
• The developed protocol relies on the partial overlap among a set of three WPs.
• Two rounds of nested PER-PCRs can generally result in a positive walking result.
Keywords: Genome walking Walking primer Partial overlap between walking primers Sequence-specific primer Partial annealing Intra-strand annealing Agarose gel electrophoresis DNA sequencing
Graphical overview
Background
In life science–related research, genome walking has achieved various applications, such as identifying transgenes, discovering new genetic resources, and cloning full-length genes, by mining unknown genomic regions flanking known DNA [2–4]. Therefore, genome walking has played an essential role in advancing life sciences–related disciplines like genetics, genomics, and microbiology [5–8].
At present, there are many genome walking methods available. The earliest genome walking method depended on constructing and then screening a genome library, which is laborious and time-consuming. Therefore, this method is seldom adopted now [9,10]. In contrast, PCR-based genome walking has received special attention due to its rapidity, low cost, and ease of operation [11–13]. To date, dozens of PCR-based genome walking methods have been established [14–17]. The existing PCR-based methods can be divided into two groups according to whether genome preprocessing is required preceding PCR: genome preprocessing-dependent PCR (such as panhandle PCR, inverse PCR, and restriction site extension PCR) and random PCR (such as wristwatch PCR, SiteFinding-PCR, and thermal asymmetric interlaced PCR). The latter has gradually become the mainstream walking method since it is free of the time-consuming genome preprocessing step. Although there are many random PCR walking methods available now, developing a practical analogous method is still popular [1,18].
Herein, a novel protocol based on primer extension refractory PCR (PER-PCR) is detailed for rapid and reliable genome walking. The PER-PCR relies on a set of three walking primers (WP): primary WP (PWP), secondary WP (SWP), and tertiary WP (TWP). The middle 15 nt of PWP overlaps the 3' part of SWP/TWP. The 5' parts of the three WPs are different from each other. The 15 nt overlap allows the WP to anneal to its predecessor site in the low annealing temperature (50 ) cycle. However, the WP complement is unable to be produced on the DNA template due to the mismatch of the two WPs 5' regions. Clearly, only target DNA is amplified in the next 65 cycles. The disadvantage of PER-PCR is the complexity of designing a practical WP set. However, the three WP sets provided in this study are universal to any species. Users need only to design the required sequence-specific primer (SSP) set. This PER-PCR protocol can be used in many aspects, such as identifying new genes, unveiling regulatory DNAs, or clarifying structures of genetic elements [1].
Materials and reagents
Biological materials
1. Genomic DNA of rice was obtained from the lab of Dr. Xiaojue Peng at Nanchang University
2. Genomic DNA of Levilactobacillus brevis CD0817 [19–22] was extracted by our lab
Reagents
1. 10× LA PCR buffer (Mg2+ plus) (Takara, catalog number: RR042A)
2. 6× Loading buffer (Takara, catalog number: 9156)
3. LA Taq polymerase (hot-start version) (Takara, catalog number: RR042A)
4. dNTP mixture (Takara, catalog number: RR042A)
5. DL 5,000 DNA marker (Takara, catalog number: 3428Q)
6. 1× TE buffer (Sangon, catalog number: B548106)
7. Agarose (Sangon, catalog number: A620014)
8. 1 M NaOH (Yuanye, catalog number: B28412)
9. 0.5 M EDTA (Solarbio, catalog number: B540625)
10. Green fluorescent nucleic acid dye (10,000 ×) (Solarbio, catalog number: G8140)
11. Tris (Solarbio, catalog number: T8060)
12. Boric acid (Solarbio, catalog number: B8110)
13. DiaSpin DNA Gel Extraction kit (Sangon, catalog number: B110092)
14. Primers (Sangon)
PWP1: AAAGTAGTCATGTATCTGCGTCCTAGTC
PWP2: AAAGTAGTCATGTATCTGGCAGTCATAG
PWP3: AAAGTAGTCATGTATCTGTGCTGTCTGA
SWP: GTCGGTCTGGGTAGTCATGTATCTG
TWP: ACCCTGTGCCGTAGTCATGTATCTG
SSP1-gadR: TCCTTCGTTCTTGATTCCATACCCT
SSP2-gadR: CCATTTCCATAGGTTGCTCCAAGGTCA
SSP3-gadR: TAGGATACTGGCTAAAATGAATTAACTCGGAT
SSP1-hyg: GGAAGTGCTTGACATTGGGGAGT
SSP2-hyg: AAGACCTGCCTGAAACCGAACTGC
SSP3-hyg: CAAGGAATCGGTCAATACACTACATGGC
Solutions
1. 2.5× TBE buffer (see Recipes)
2. 0.5× TBE buffer (see Recipes)
3. 100 μM primer (see Recipes)
4. 10 μM primer (see Recipes)
5. 1.5% agarose gel (see Recipes)
Recipes
1. 2.5 × TBE buffer
Reagent Final concentration Amount
0.5 M EDTA solution 5 mM 10 mL
Tris 225 mM 27 g
Boric acid 225 mM 13.75 g
ddH2O n/a 980 mL
Total n/a 1,000 mL
Adjust pH to 8.3 with 1 M NaOH and then replenish the solution to 1,000 mL with ddH2O.
2. 0.5× TBE buffer
Reagent Final concentration Amount
2.5× TBE buffer 0.5× 200 mL
ddH2O n/a 800 mL
Total n/a
3. 100 μM primer
Reagent Final concentration Quantity or Volume
Powdery primer 100 μM n/a
1× TE buffer 1× Volume specified in the sheet of primer synthesis
Total n/a Volume specified in the sheet of primer synthesis
Note: Dilute a portion of the 100 μM primer to prepare 10 μM primer and store the remaining portion at -80 .
4. 10 μM primer
Reagent Final concentration Quantity or Volume
100 μM primer 10 μM 1 μL
1× TE buffer 1× 9 μL
Total n/a 10 μL
Note: Prepare extra volume of a 10 μM primer and divide it into multiple 1.5 mL microcentrifuge tubes; then, store the microcentrifuge tubes at -80 . Take one tube at a time and store it at -20 after use.
5. 1.5% agarose gel
Reagent Final concentration Quantity or Volume
Agarose 1.5% 1.5 g
0.5× TBE buffer 0.5× 100 mL
Green fluorescent nucleic acid dye (10,000 ×) 1× 10 μL
Total n/a 100 mL
Laboratory supplies
1. 0.2 mL thin-wall PCR tubes (Kirgen, catalog number: KG2311)
2. 10 μL pipette tips (Sangon, catalog number: F600215)
3. 200 μL pipette tips (Sangon, catalog number: F600227)
4. 1,000 μL pipette tips (Sangon, catalog number: F630101)
5. 1.5 mL microcentrifuge tubes (Labselect, catalog number: MCT-001-150)
Equipment
1. PCR apparatus (Analtytikjena, model: Biometra TAdvanced)
2. Electrophoresis apparatus (Beijing Liuyi, model: DYY-6C)
3. Gel imaging system (Bio-Rad, model: ChemiDoc XRS+)
4. Microcentrifuge (Tiangen, model: TGear)
Software and datasets
1. Oligo 7 software (Molecular Biology Insights, Inc., USA)
2. DNASTAR Lasergene software (DNASTAR, Inc., USA)
Procedure
The overview of PER-PCR is shown in Figure 1.
Figure 1. Schematic overview of PER-PCR. PWP: primary walking primer; SWP: secondary walking primer; TWP: tertiary walking primer; SSP: sequence-specific primer. Thin solid line: known sequence; thin dotted line: unknown sequence; arrows: primers; thick lines: primer complements; symbol +: plus strand; and symbol -: minus strand.
Note: In primary PER-PCR, the initial 65 °C cycles only permit SSP1 to hybridize with its complement on known region, thus producing several target plus strands (+). In the subsequent 25 °C cycle, PWP randomly anneals somewhere on the unknown region of this strand and synthesizes the target minus strand (-). In the following 65 °C cycles, this target minus strand is exponentially amplified. Any non-target single strand formed in the 25 °C cycle cannot be amplified because it lacks an authentic binding site for any primer. In secondary PCR, the three types (type I, primed by SSP1; type II, primed by SSP1 and PWP; and type III, primed by PWP) of primary PCR non-target products are easily removed, as they lack an exact binding site for at least one of the primers. Nevertheless, the target product is exponentially amplified. Similarly, only the target product is further amplified in tertiary PCR.
A. Design of primers
1. Design WP sets (Figure 2).
Figure 2. The three WP sets designed in this study. The three PWPs (PWP1, PWP2, and PWP3) have a length of 28 nt, comprising three regions. Region 1 is the 5' head of three adenosines (AAA); region 2 is the 15 nt overlap in the middle; and region 3 is the 3' tails (10 nt) different from each other. SWP or TWP has a length of 25 nt, with the 15 nt overlap at the 3' part. The 5' parts of SWP and TWP are different from each other and also different from region 1 of PWP. PWP: primary walking primer, SWP: secondary walking primer, and TWP: tertiary walking primer.
Note: The DNA sequences of three WPs (PWP, SWP, and TWP) in a WP set are arbitrary, but any WP should a) have a Tm value between 60 and 65 ; b) have an even distribution of G, C, T, and A, with a GC content of about 50%; and c) avoid a hairpin or dimer structure with a Tm value exceeding 40 . The Tm value of the 15 nt overlap is about 35 °C. Simultaneously designing more than one WP set can perform parallel PER-PCRs in a single walking cycle. n WP sets comprise n PWPs, one SWP, and one TWP. The SWP and WTP are universal to the n PWPs. In general, 2–3 parallel PER-PCR sets are suggested for a single walking cycle.
a. Open the Oligo 7 software, then click File and New Sequence (Figure 3a); type in a 28 nt arbitrary sequence as the initial PWP1 in the Edit Sequence dialog box (Figure 3b). Next, click Accept/Discard and Accept (Figure 3c).
Note: Here, the design of WP set 1 (including PWP1, SWP, and TWP) is provided as an example.
Figure 3. Screenshots displaying how to enter the initial primary walking primer in the Oligo 7 software. Locations of New Sequence (a), Edit Sequence dialog box (b), and Accept (c).
b. Click Analyze, Duplex Formation, and Current Oligo (Figure 4a) to analyze primer dimer (Figure 4b).
Figure 4. Screenshots displaying how to analyze primer dimer. (a) Locations of Duplex Formation and Current Oligo. (b) Predicted primer dimers.
c. Click Analyze, Hairpin Formation, and Current Oligo (Figure 5a) to analyze primer hairpin (Figure 5b).
Figure 5. Screenshots displaying how to analyze primer hairpin. (a) Locations of Hairpin Formation and Current Oligo. (b) Predicted primer hairpins.
Note: Optimize the sequence of the initial PWP1 (see below) if it is unsatisfactory because it forms an obvious primer dimer(s) or hairpin(s).
d. Click Edit and Entire Sequence (Figure 6a) to return to the Edit Sequence dialog box (Figure 3b); edit the initial sequence according to the above analysis results, then click Accept/Discard and Accept (Figure 3c). Next, minimize this dialog box to display the dialog box shown in Figure 4a.
Figure 6. Screenshots displaying how to optimize the sequence of the initial PWP1. (a) Screenshot displaying how to return to the Edit Sequence dialog box. (b) Predicted primer hairpins.
e. Repeat steps A1b,c to evaluate the formation of primer dimer and hairpin in the edited PWP1 until a satisfactory PWP1 (Figure 6b) is obtained.
Note: The PWP1 shown in Figure 6b is satisfactory because this oligo only forms an acceptable primer dimer and does not form any hairpin.
f. Select region 2 (the middle 15 nt overlap) of the satisfactory PWP1.
g. Attach a 10 nt arbitrary oligo to the 5’ end of this overlap, creating the initial SWP.
h. Assess this SWP using the above method (corresponding to the steps A1a–e) until a satisfactory SWP is obtained.
i. Design a satisfactory TWP like designing the SWP.
Critical: An obvious primer dimer should be evaded between any two WPs in a WP set.
2. Select an SSP set of three SSPs (outmost SSP1, middle SSP2, and innermost SSP3) from a known DNA (Figure 7).
Figure 7. Mutual positional relationship of SSP1, SSP2, and SSP3. SSP: sequence-specific primer; thin solid line: known sequence; thin dotted line: unknown sequence; arrows: primers; thick lines: primer complements.
Note: An SSP set of three SSPs is within the same known DNA, with an extension direction toward the unknown flanking region. An SSP should (i) have a length of approximately 25 nt, with a melting temperature (Tm) between 60 and 65 ; (ii) have an even distribution of guanine (G), cytosine (C), thymine (T), and adenine (A), with a GC content of about 50%; (iii) avoid repetition sequences; and (iv) avoid a hairpin or dimer structure with a Tm exceeding 40 . In general, there is no limit of distance between the two adjacent SSPs, but a distance of approximately 20–50 nt is suggested if possible.
j. Open the Oligo 7 software, then click File and Open to enter the known DNA sequence of the gadR gene (Figure 8a).
Note: Here, the design of SSP1-gadR is provided as an example.
Figure 8. Screenshots displaying how to design SSP1-gadR. (a) Screenshot displaying how to enter the known DNA sequence. (b) Screenshot displaying how to choose an oligo with a defined length. (c) The predicted primer dimers and hairpin.
k. Click Change and Current Oligo Length to define the length (for instance 25 nt) of oligo to be analyzed; then, move the mouse to a position of the entered sequence (the 25 nt following the mouse is automatically shaded) and click to select this shaded 25 nt (Figure 8b).
l. Assess the selected 25 nt oligo according to the method (corresponding to the steps of Figures 4 and 5) for assessing PWP1.
Note: In case the current SSP1-gadR is unsatisfactory due to forming an obvious primer dimer(s) or hairpin(s), re-select and assess a new oligo by repeating steps A1a–c until a satisfactory SSP1-gadR (Figure 8c) is obtained.
m. Design the nested SSP2-gadR and SSP2-gadR, just like designing the SSP1-gadR.
Critical: According to Mazars et al. [23], each SSP shows a melting temperature of 65–70 °C and should avoid forming severe hairpin or dimer structure.
B. PER-PCR amplification
The current protocol comprises three rounds of nested PCRs. Primary PCR is driven by PWP and SSP1; secondary PCR is driven by SWP and SSP2; and tertiary PCR is driven by TWP and SSP3.
1. Primary PER-PCR
a. Pipette primary PER-PCR components (Table 1) into a 0.2 mL PCR tube.
Table 1. Primary PER-PCR mix
Reagent Final concentration Amount (μL)
Genomic DNA Microbe, 0.2–2 ng/μL; plant or animal, 2–20 ng/μL 1
LA Taq polymerase (5 U/μL) 0.05 U/μL 0.5
PWP (10 μM) 0.2 μM 1
SSP1 (10 μM) 0.2 μM 1
10× LA PCR buffer II (Mg2+ plus) 1× 5
dNTP mixture (2.5 mM each) 0.4 mM each 8
ddH2O n/a 33.5
Total n/a 50
b. Completely mix the components with a pipette.
c. Centrifuge for 10–20 s with a microcentrifuge to gather the mixture.
d. Run the amplification in the PCR apparatus (Table 2).
Table 2. Primary PER-PCR cycling conditions
Step Temp. (°C) Duration Cycle
Initial denaturation 95 1 min 1
Denaturation 95 20 s 5
Annealing 65 30 s
Extension 72 2 min
Denaturation 95 20 s 1
Annealing 25 30 s
Extension 72 2 min
Denaturation 95 20 s 30
Annealing 65 30 s
Extension 72 2 min
Final extension 72 5 min 1
Hold 4 forever
e. Put the PCR product onto ice.
f. Take 1 μL of the product as the template of secondary PER-PCR.
g. Store the remaining product at -20 °C for future assays.
2. Secondary PER-PCR
a. Pipette secondary PER-PCR components (Table 3) into a 0.2 mL PCR tube.
Table 3. Secondary PER-PCR mix
Reagent Final concentration Amount (μL)
Primary PCR product n/a 1
LA Taq polymerase (5 U/μL) 0.05 U/μL 0.5
SWP (10 μM) 0.2 μM 1
SSP2 (10 μM) 0.2 μM 1
10× LA PCR buffer II (Mg2+ plus) 1× 5
dNTP mixture (2.5 mM each) 0.4 mM each 8
ddH2O n/a 33.5
Total n/a 50
Critical: Dilute primary PER-PCR product 10–10,000 folds if necessary.
b. Completely mix the components with a pipette.
c. Centrifuge for 10–20 s with a microcentrifuge to gather the mixture.
d. Run the amplification in the PCR apparatus (Table 4).
Table 4. Secondary PER-PCR cycling conditions
Step Temp. (°C) Duration Cycle
Initial denaturation 95 1 min 1
Denaturation 95 20 s 1
Annealing 50 30 s
Extension 72 2 min
Denaturation 95 20 s 25
Annealing 65 30 s
Extension 72 2 min
Final extension 72 5 min 1
Hold 4 forever
e. Put the PCR product onto ice.
f. Take 1 μL of the product as the template of tertiary PER-PCR.
g. Store the remaining product at -20 °C for future assays.
3. Tertiary PER-PCR
a. Pipette tertiary PER-PCR components (Table 5) into a 0.2 mL PCR tube.
Table 5. Tertiary PER-PCR mix
Reagent Final concentration Amount (μL)
Secondary PCR product n/a 1
LA Taq polymerase (5 U/μL) 0.05 U/μL 0.5
TWP (10 μM) 0.2 μM 1
SSP3 (10 μM) 0.2 μM 1
10× LA PCR buffer II (Mg2+ plus) 1× 5
dNTP mixture (2.5 mM each) 0.4 mM each 8
ddH2O n/a 33.5
Total n/a 50
Critical: Dilute secondary PER-PCR product 10–10,000 folds if necessary.
b. Completely mix the components with a pipette.
c. Centrifuge for 10–20 s with a microcentrifuge to gather the mixture.
d. Run the amplification in the PCR apparatus (Table 6).
Table 6. Tertiary fork PCR cycling conditions
Step Temp. (°C) Duration Cycle
Initial denaturation 95 1 min 1
Denaturation 95 20 s 1
Annealing 50 30 s
Extension 72 2 min
Denaturation 95 20 s 25
Annealing 65 30 s
Extension 72 2 min
Final extension 72 5 min 1
Hold 4 forever
e. Store the PCR product at -20 °C for future assays.
C. Gel electrophoresis
1. Completely mix 5 μL of PER-PCR product and 1 μL of 6× loading buffer.
2. Load the mixture into a 1.5% agarose gel supplemented with 1× green fluorescent nucleic acid dye.
3. Set the electrophoresis apparatus to a voltage of 150 V (the distance between the two electrodes is 30 cm).
4. Check the PCR product using the ChemiDoc XRS+ imaging system after approximately 25 min of electrophoresis (Figure 9).
Figure 9. Mining unknown flanks of gadR (A) and hyg (B). The three parallel sets of PER-PCRs in each walking are shown. Lanes P1, P2, and P3: primary products; Lanes S1, S2, and S3: secondary products; Lanes T1, T2, and T3: tertiary products; and Lanes Ms: DNA5000 Marker (5,000, 3,000, 2,000, 1,500, 1,000, 750, 500, 250, and 100 bp). The clear PCR amplicons are shown with white arrows.
D. Recovery of PCR product
1. Completely mix 40 μL of secondary/tertiary PER-PCR product and 8 μL of 6× loading buffer.
2. Load the mixture into a 1.5% agarose gel supplemented with 1× green fluorescent nucleic acid dye.
3. Set the electrophoresis apparatus to a voltage of 150 V (the distance between the two electrodes is 30 cm).
4. Visualize the PCR product using the ChemiDoc XRS+ imaging system. Subsequently, cut out clear DNA band(s) using a knife.
5. Extract the DNA band(s) from the cut gel using the DiaSpin DNA Gel Extraction kit.
6. Confirm the extracted DNA band(s) with 1.5% agarose gel electrophoresis.
E. DNA Sequencing
Sequence the extracted DNA band(s) at Sangon Biotech Co., Ltd.
Data analysis
Perform sequence analysis using the By Clustal W Method function in MegAlign software. In general, walking is considered correct if one end of the obtained DNA overlaps with one end of the known DNA [24–26].
1. Open the MegAlign software, then click File and Enter Sequences (Figure 10a) to input a PER-PCR product and the corresponding known DNA sequence (Figure 10b).
Figure 10. Screenshots displaying how to enter DNA sequences. (a) Screenshot displaying the location of Enter Sequences. (b) Input DNA sequences.
2. Click Align and By Clustal W Method (Figure 11a) to output the alignment outcome (Figure 11b).
Note: Genome walking is regarded as successful if the SSP3-sided region of the PER-PCR product overlaps the known DNA (an example is shown in Figure 11b).
Figure 11. Screenshots displaying how to align the entered sequences. (a) Screenshot showing the location of By Clustal W. (b) Alignment outcome.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article:
Li et al. [1] Primer extension refractory PCR: an efficient and reliable genome walking method. Molecular Genetics and Genomics (Figure 9).
General notes and troubleshooting
General notes
1. PER-PCR comprises three rounds of nested PCRs. However, secondary PCR generally suffices to release a positive outcome.
2. In general, it is not necessary to check the primary PER-PCR product.
3. Before tertiary PER-PCR, secondary PER-PCR product is analyzed using agarose gel electrophoresis so as to see if a clear DNA band(s) appears. If a clear DNA band(s) appears, it is recovered and sequenced. It is not necessary to perform tertiary PCR if this DNA band(s) is the target product.
4. Like the other existing PCR-based genome walking protocols, the current PER-PCR protocol also suffers from the multiple-band issue. However, if multiple DNA bands appear, only the largest one needs to be assayed.
5. Simultaneously design n WP sets (2–3 sets are suggested) so as to do n parallel PER-PCRs in a single walking cycle. The parallel PER-PCR design of this protocol is conducive to improving the success and efficiency of genome walking. n sets of WPs comprise n PWPs, one SWP, and one TWP. The SWP and WTP are universal to the n PWPs.
6. The current PER-PCR protocol is applicable to any species.
Troubleshooting
Problem 1: There is no clear DNA band(s) in secondary or even tertiary PER-PCR.
Possible cause: (i) In primary PER-PCR, target amplification is weak while non-target amplification is strong. (ii) the GC content of the genomic DNA template may be high.
Solution: (i) Dilute primary PER-PCR product 10–10,000 times and then use 1 μL of each dilution as the template in secondary PER-PCR. Next, tertiary PER-PCR is performed using each secondary PER-PCR product as the template. If there is still no clear DNA(s) in any secondary/tertiary PER-PCR, redesign an SSP set. (ii) Add a PCR enhancer (for example betaine) to PCR reaction solution, increase the working concentration of primers, or lengthen the 5’ head of walking primer to enhance its Tm value; meanwhile, use an SSP with a higher Tm value.
Problem 2: A PER-PCR product cannot be directly sequenced.
Possible cause: There is interference from non-target products.
Solution: Clone the obtained DNA band and then sequence.
Problem 3: A clear DNA band(s) is the non-target product.
Possible cause: The genome has sites homologous to the SSP complements in other region(s).
Solution: Redesign an SSP set at different sites of known DNA.
Problem 4: The PCR product cannot be separated by agarose gel electrophoresis, showing as a large block in the gel.
Possible cause: The walking primer has multiple major partial annealing sites on unknown flanks that are very close to each other.
Solution: Recover the upper part of this block from the gel and then directly use SSP3 or an SSP inner to SSP3 to sequence it or clone it and then sequence.
Acknowledgments
This work was funded by the Jiangxi Provincial Department of Science and Technology (grant No. 20225BCJ22023), China, and the National Natural Science Foundation of China (grant No 32160014). This PER-PCR-based genome walking protocol has been originally described and validated in Molecular Genetics and Genomics [1].
Competing interests
The authors declare no competing interests.
References
Li, H., Lin, Z., Guo, X., Pan, Z., Pan, H. and Wang, D. (2024). Primer extension refractory PCR: an efficient and reliable genome walking method. Mol Genet Genomics. 299(1): 27. https://doi.org/10.1007/s00438-024-02126-5
Thirulogachandar, V., Pandey, P., Vaishnavi, C. and Reddy, M. K. (2011). An affinity-based genome walking method to find transgene integration loci in transgenic genome. Anal Biochem. 416(2): 196–201. https://doi.org/10.1016/j.ab.2011.05.021
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Lin, Z., Wei, C., Pei, J. and Li, H. (2023). Bridging PCR: An efficient and reliable scheme implemented for genome-walking. Curr Issues Mol Biol. 45(1): 501–511. https://doi.org/10.3390/cimb45010033
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Chang, K., Wang, Q., Shi, X., Wang, S., Wu, H., Nie, L. and Li, H. (2018). Stepwise partially overlapping primer-based PCR for genome walking. AMB Express. 8(1): 77. https://doi.org/10.1186/s13568-018-0610-7
Wei, C., Lin, Z., Pei, J., Pan, H. and Li, H. (2023). Semi-site-specific primer PCR: A simple but reliable genome-walking tool. Curr Issues Mol Biol. 45(1): 512–523. https://doi.org/10.3390/cimb45010034
Chen, H., Wei, C., Lin, Z., Pei, J., Pan, H. and Li, H. (2024). Protocol to retrieve unknown flanking DNA sequences using semi-site-specific PCR-based genome walking. STAR Protoc. 5(1): 102864. https://doi.org/10.1016/j.xpro.2024.102864
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Alquezar‐Planas, D. E., Löber, U., Cui, P., Quedenau, C., Chen, W. and Greenwood, A. D. (2020). DNA sonication inverse PCR for genome scale analysis of uncharacterized flanking sequences. Methods Ecol Evol. 12(1): 182–195. https://doi.org/10.1111/2041-210x.13497
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Li, H., Sun, T., Jia, M., Wang, L., Wei, C., Pei, J., Lin, Z. and Wang, S. (2022). Production of gamma-aminobutyric acid by Levilactobacillus brevis CD0817 by coupling fermentation with self-Buffered whole-cell catalysis. Fermentation 8(7): 321. https://doi.org/10.3390/fermentation8070321
Jia, M., Zhu, Y., Wang, L., Sun, T., Pan, H. and Li, H. (2022). pH auto-sustain-based fermentation supports efficient gamma-aminobutyric acid production by Lactobacillus brevis CD0817. Fermentation.. 8(5): 208. https://doi.org/10.3390/fermentation8050208
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Article Information
Publication history
Received: Sep 14, 2024
Accepted: Dec 12, 2024
Available online: Dec 26, 2024
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Molecular Biology > DNA > PCR
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5,189 | https://bio-protocol.org/en/bpdetail?id=5189&type=0 | # Bio-Protocol Content
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Peer-reviewed
Novel Workflows for Separate Isolation of Pathogen RNA or DNA from Wastewater: Detection by Innovative and Conventional qPCR
KB Kristina M. Babler *
HS Helena M. Solo-Gabriele *
MS Mark E. Sharkey *
AA Ayaaz Amirali *
(*contributed equally to this work)
In Press, Available online: Jan 14, 2025
DOI: 10.21769/BioProtoc.5189 Views: 42
Reviewed by: Lucy XieSonali ChaturvediWeidong An
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Original Research Article:
The authors used this protocol in Journal of Biomolecular Techniques Dec 2023
Abstract
Wastewater-based surveillance (WBS) can provide a wealth of information regarding the health status of communities from measurements of nucleic acids found in wastewater. Processing workflows for WBS typically include sample collection, a primary concentration step, and lysis of the microbes to release nucleic acids, followed by nucleic acid purification and molecular-based quantification. This manuscript provides workflows from beginning to end with an emphasis on filtration-based concentration approaches coupled with specific lysis and nucleic acid extraction processes. Here, two WBS processing approaches are presented, one focusing on RNA-specific pathogens and the other focused on DNA-specific pathogens found within wastewater: 1) The RNA-specific approach, employed for analyzing RNA viruses like severe acute respiratory syndrome coronavirus-2 (SARS-CoV-2) couples electronegative filtration of wastewater with the placement of the filter within a lysis buffer followed by direct RNA extraction. 2) The DNA-specific approach, employed for analyzing DNA pathogens like Candida auris, uses size selection membranes during filtration, subsequently followed by a lysis buffer, bead-beating, and DNA extraction. Separate workflows for RNA versus DNA isolations have the advantage of improving the detection of the target pathogen. A novel aspect of the RNA-specific workflow is the direct extraction of nucleic acids from filter lysates, which shows enhanced recoveries, whereas the DNA-specific approach requires bead beating prior to extraction. Novelty is also provided in a new qPCR approach called Volcano 2nd Generation (V2G), which uses a polymerase capable of using RNA as a template, bypassing the reverse transcriptase step normally required for qPCR.
Key features
• Membrane filtration approaches for concentrating suspended solids from wastewater. After concentration, workflows are optimized for separate recovery of RNA and DNA.
• Unique polymerase utilized to perform qPCR analysis, foregoing reverse transcription, for RNA.
• Sample products for use with other molecular techniques (e.g., sequencing) as workflow approaches generate high-quality, concentrated nucleic acid extracts with minimal inhibitors.
• Validated through COVID-19 surveillance where >1,000 samples of wastewater and >3,000 filter concentrates produced from these samples have been created and analyzed, with published results.
Keywords: Wastewater Pathogen detection Nucleic acids Bead beating Membrane filtration
Graphical overview
Laboratory workflow of sample preparation
Background
The use of wastewater for surveying the health of communities has been a growing application in the field of public health. Analyzing environmental samples like wastewater can inform researchers of current infectious disease spread or the general health of communities. More specifically, wastewater can capture spatial and temporal trends of pollutants, drug abuse in communities, and antimicrobial resistance genes [1]. Recent wastewater-based surveillance (WBS) literature has emphasized not only WBS’s rapid development but also shed light on the importance of method selection prior to experimentation. The paper by Pecson et al. [2] is a primary example of such literature, where 32 participating laboratories at the start of the COVID-19 pandemic cooperated to compare strategies for pathogen surveillance from wastewater. Their focus was on reproducibility, sensitivity, and the impact that “other method steps” have on the recovery of viruses like SARS-CoV-2. Since the publication of Pecson et al. [2], many researchers have initiated WBS programs and have focused on comparing specific methods; a general theme has been to expand WBS’s application for SARS-CoV-2 detection to other pathogens of interest to expand WBS as a tool for public health [3–11,12,13].
Although methods are merging toward the isolation of total nucleic acids (RNA and DNA simultaneously) using the same extraction procedure, studies have shown [2] that optimal recoveries require different extraction procedures. Here, we propose two different concentration-plus-extraction strategies—one for the recovery of RNA and another for the recovery of DNA, both optimized for wastewater. The assessment of samples with the Volcano 2nd Generation (V2G) qPCR assay adds an additional uniqueness to this procedure due to the equal performance of V2G to conventional RT-qPCR [6] while providing a decrease in sample analysis time. While filtration-based concentration is not a novel concept in and of itself, its use, coupled with sample type, membrane selection, filtration protocol, lysis method, and downstream molecular techniques, is unique and adjustable for specific research needs. For example, varying volumes of wastewater can be used depending on the composition of each sample (turbidity, suspended solids concentration, etc.). Additionally, different membrane types and pore sizes can be utilized to fit the specific application. The filtration approach can be optimized for the capture of viruses by charge or for larger microbes by size exclusion. This includes pathogens like Candida auris, a growing health concern in many communities [14–16]. The use of bead beating, especially coupled with filtration approaches, is also advantageous as it increases the likelihood of detecting pathogens with cell membranes like C. auris [16] from wastewater. However, bead beating is not recommended for the recovery of RNA viruses. Bead beating can limit the detection of RNA viruses due to heat and mechanical agitation, which can lead to the degradation of some nucleic acids if not optimized prior to experimentation [3]; it also extracts ribosomal RNA from bacteria and higher forms of microbes, confounding the RNA signal from viruses. The inclusion of chemicals like HCl for the recovery of viruses by charge, although recommended in this protocol, could also impact recovery, as seen in Babler et al. [3]. Regarding limitations, filtration-based workflows require sterilization of equipment unless disposable options are favored, so they can be costly to initiate. Moreover, the processes presented here from start to finish are designed to be completed manually as opposed to magnetic bead concentration using robotic approaches [18], so laboratory capacity and personnel bandwidth also play into the ability to use these methods if many samples are expected to be analyzed. To expand these protocols to evaluate pathogens beyond SARS-CoV-2 and C. auris, optimization is recommended to ensure that recoveries of the intended pathogen are suitable for downstream applications. These methods, as they have been optimized for wastewater, may not be generalizable for other sample types.
Materials and reagents
Reagents
Electronegative filtration and RNA extraction workflow: specific materials and reagents
1. Hydrochloric acid 10% (Spectrum Chemical MFG Corp., catalog number: HY105)
2. pH standards 4, 7, 10 (Cole-Parmer, catalog number: UX-05942-10)
3. Magnesium chloride, MgCl2, 51% w/v aqueous solution (RICCA Chemical Company, catalog number: 4470)
4. Zymo QuickRNA Viral Kit (Zymo Research, catalog number: R1034/1035)
5. V2G buffer (MyPols, catalog number: 8100)
6. V2G polymerase (MyPols, catalog number: 8400M)
7. Rox (Thermo Fisher Scientific, catalog number: 12223012)
8. dNTPs (Thermo Fisher Scientific, catalog number: R0193)
9. Platinum Taq Antibody (TaKaRa, catalog number: 9002A)
Vacuum filtration and DNA extraction workflow: specific materials and reagents
1. ZymoBIOMICS DNA Miniprep kit (Zymo Research, catalog number: D4300)
2. TaqMan Fast Universal PCR master mix (2×) (Thermo Fisher, catalog number: 4352042)
Materials and reagents used for both workflows
1. Vero Cells [American Type Culture Collection (ATCC), catalog number: CCL-81]
2. Luria Bertani (LB) Broth (Thermo Fisher Scientific, catalog number: 10855001)
3. LB agar plates (Thermo Fisher Scientific, catalog number: 22700025)
4. Tween 80 (Millipore Sigma, catalog number: P1754-25ML)
5. 100 mg/mL ampicillin (Millipore Sigma, catalog number: A5354-10ML)
6. Mycobacterium smegmatis mc2155 (NIH AIDS Reagent Program, catalog number: 2195)
7. Beta coronavirus 1, strain OC43 (ATCC, catalog number: VR-1558)
8. RPMI media (Thermo Fisher Scientific, catalog number: 12633012)
9. Wastewater [collected in pre-labeled pre-sterilized bottles (500 mL capacity) containing 0.5 mL of sterile 100 g/L sodium thiosulfate; details of bottles and reagents below]
10. 1× DNA/RNA shield (Zymo Research, catalog number: R1100-250)
11. OneStep PCR Inhibitor Removal kit (Zymo Research, catalog number: D6030)
12. qPCR primers (synthesized by Sigma-Aldrich)
13. qPCR probes (synthesized by Integrated DNA Technologies)
14. Nuclease-free water (Fisher Scientific, catalog number: AM9930)
15. 99.5% isopropanol (Thermo Fisher Scientific, catalog number: 149320050)
16. 70% ethanol (Fisher Scientific, catalog number: BP82031GAL)
17. 100% ethanol (Fisher Scientific, catalog number: BP2818500)
18. Sodium thiosulfate (GrowingLabs, catalog number: LC250001)
19. Clorox bleach (concentrated)
Laboratory supplies
Electronegative filtration and RNA extraction workflow: specific laboratory supplies
1. MCE membrane filter, 47 mm, 0.45 μm pore size (Millipore Sigma, catalog number: HAWP04700)
2. Magnetic stir bars [12 mm (L) × 3 mm (D)] (VWR, catalog number: 58948-091)
3. DI water (Thermo Fisher Scientific, catalog number: 15230001)
4. Glass dropper (Fisher Scientific, catalog number: 14-955-502)
5. 5 mL Eppendorf Tubes (sterile) (Eppendorf, catalog number: 0030119460)
Vacuum filtration and DNA extraction workflow: specific laboratory supplies
1. Pall GN-6 Metricel MCE membrane disc filters, 47 mm, 0.45 μm pore size (New Star Environmental, catalog number: 66278)
2. 2 mL ZR BashingBead lysis tubes (0.1 & 0.5 mm) (Zymo Research, catalog number: S6012-50)
3. Polystyrene, disposable, sterile Petri dishes (Carolina, catalog number: 741248)
4. Disposable sterile scalpels #10 (Surgical Supply Service, catalog number: 75745)
Laboratory supplies used for both workflows
1. Biohazard waste benchtop container (1.4 L) (VWR, catalog number: 11214-708)
2. Vactrap 2 vacuum trap system for aspiration and vacuum protection (VWR, catalog number: 76207-602)
3. Pall magnetic filter funnels, 47 mm [Weber Scientific, catalog number: EF8452B (300 mL), EF8452F (500 mL)]
4. Clamps and chains to hold sample collection bottles used to collect samples from sewer holes in the field [available from any hardware store. Hose clamps large enough to attached to bottles (MIAHART, 6-inch hose clamp adjustable 304 stainless steel duct clamps worm gear), 1/8-inch chain hooks with locking connector (BNYZWOT, D-shape, twist lock), and lightweight utility chain are recommended (4every SUS304). The chain hooks are used to connect the hose clamps holding the bottles to two chains, one on each side of the bottle]
5. Strainer/colander for field (Bradshaw Home, catalog number: 72115); the strainer should have 2–3 mm diameter openings, kitchen type strainer will work
6. Funnel for field (Walmart, catalog number: HTFF-2020); the funnel should be wide at the top and narrow enough at the bottom to fit easily into the sample collection bottles
7. 5-gallon heavy-duty plastic buckets to capture spillage when pouring wastewater samples (Lowe’s, catalog number: 954434)
8. 2 L sterile plastic wide-mouth collection bottles (Thermo Fisher Scientific, catalog number: 2120-0005 if disposable, or Thermo Fisher Scientific, catalog number: 2121-0005 for reuse via autoclaving)
9. 500 mL sterile plastic collection bottles (VWR, catalog number: 16060-012 if disposable, or VWR, catalog number: 16060-012 for reuse via autoclaving); a 500 mL volume line marked on the bottle using a paint pen
10. 1 L sterile wide-mouth collection bottles (VWR, catalog number: 16060-014 if disposable, or VWR, catalog number: 16060-014 for reuse via autoclaving)
11. Kimwipes (Grainger, catalog number: 36VC06)
12. Graduated cylinders made of polypropylene for reuse by autoclaving (100 mL) (Carolina, catalog number: 721603)
13. Spade smooth tip forceps (AmScope, catalog number: TW-487)
14. PYREX griffin beakers (100 mL) (Millipore Sigma, catalog number: CLS1000100-12EA)
15. P5000 pipette (Gilson, catalog number: F144066)
16. P5000 pipette tips (Thermo Fisher Scientific, catalog number: 94052550)
17. P1000 pipette (Gilson, catalog number: F144059M)
18. P1000 filtered pipette tips (IBIS Scientific, catalog number: M-1000-9FC)
19. P200 Pipette (Gilson, catalog number: F144058M)
20. P200 filtered pipette tips (IBIS Scientific, catalog number: M-0200-9FC)
21. P100 pipette (Gilson, catalog number: F144057M)
22. P100 filtered pipette tips (IBIS Scientific, catalog number: M-0100-9FC)
23. P20 pipette (Gilson, catalog number: F144056M)
24. P20 filtered pipette tips (IBIS Scientific, catalog number: M-0020-9FC)
25. P10 pipette (Gilson, catalog number: F144055M)
26. P10 pipette tips (IBIS Scientific, catalog number: M-0010-9FC)
27. Tube racks (Eppendorf, 1.5 mL and 2 mL, catalog number: 0030119819; 5 mL and 15 mL, catalog number: 0030119827)
28. Autoclave bins (Fisher Scientific, catalog number: 13-359-20B)
29. T175 cell culture flasks (Fisher Scientific, catalog number: 10-126-34)
30. Serological pipettor (Pipette.com, catalog number: DP-501R)
31. 10 mL serological pipettes (Fisher Scientific, catalog number: NC9868325)
32. 50 mL conical tubes (Fisher Scientific, catalog number: 14-432-22)
33. 96-well plates (Bio-Rad, catalog number: HSP9601)
34. Bio-Rad microseal “B” seal (Bio-Rad, catalog number: MSB1001)
35. 1.5 mL microcentrifuge tubes (Fisher Scientific, catalog number: 05-408-129)
36. Labels for all tubes (Brady, catalog number: THT-59-7425-2-SC)
37. Freezer storage boxes (81 slot: Alkali Scientific, catalog number: FB2CC-81; 25 slot: Eppendorf, catalog number: 0030140532)
38. Laboratory coats (Fisher Scientific, catalog number: 19-181-594)
39. Disposable laboratory coats for field (Dupont Tyvek 400, catalog number: TY212S)
40. Gloves (VWR, catalog number: 76582-340)
41. Eye protection (Fisher Scientific, catalog number: 19-039-590)
42. (Optional) respiratory protection (Breatheze KN95 disposable face masks, Amazon)
43. Laboratory data sheets on clipboard with pen (created depending on lab needs using Microsoft Word or Excel)
44. Permanent paint pen (ULINE, catalog number: S-20622)
Equipment
Electronegative filtration and RNA extraction workflow: specific equipment
1. pH Meter (Orion Star) (Thermo Fisher Scientific, catalog number: STARA2110)
2. Magnetic stir plate (GrowingLabs, catalog number: H4000-S)
3. Vortex Genie 2 (Scientific Industries, Inc., catalog number: SI-0236)
Vacuum filtration and DNA extraction workflow: specific equipment
1. BeadRuptor 12 Instrument (Omni International)
Equipment used for both workflows
1. YSI probe (Xylem YSI ProDSS, catalog number: 626870-1); for measuring basic water quality of samples. Recommend adding probes for pH (#626904), water temperature and salinity (#626902), turbidity (#626901), and dissolved oxygen (#626900)
2. Clean water reservoir with hand pump to rinse of equipment after sample collection (Itisll, portable garden pump sprayer, Size: 2 Gal_Brasswand, ULINE, catalog number: S-20860)
3. (Optional) Refrigerated autosampler (HACH, catalog number: AS950 fitted with an IO9000 for flow proportional sampling) or non-refrigerated autosampler (Teledyne ISCO, catalog number: ISCO 6712, fitted with a 2150 area velocity meter for flow paced sampling; otherwise, use time paced). Refrigerated with flow measurement preferred
4. BSL-2 hood
5. Whatman vacuum manifold (LabFilterz, catalog number: 10498761)
6. MyFuge centrifuge (5 mL capacity) (Grayline Medical, catalog number: C1005)
7. Eppendorf centrifuge (2 mL capacity) (Eppendorf, catalog number: 5420000245)
8. Benchtop centrifuge (Southern Labware, catalog number: C1008-B)
9. Bio-Rad CFX connect RealTime qPCR instrument (Bio-Rad Laboratories, catalog number: 1855201)
10. Plate spinner (GrowingLabs, catalog number: C2000-115V)
11. Synergy BioTek plate reader (Fisher Scientific, catalog number: BTS1LASI)
12. 37 °C shaker/incubator (Fisher Scientific, catalog number: 50-195-3922)
13. Spectrophotometer (Eppendorf, model: 6135)
14. Culture plate rocker (Millipore Sigma, catalog number: Z742529-1EA)
15. Corning LSE digital dry bath heater (Fisher Scientific, catalog number: 07-203-024)
16. Igloo or Coleman cooler (purchased from any general store)
17. Access to 4 °C, -20 °C, and -80 °C refrigerators and freezers. Freezers should not include defrost cycles to facilitate the preservation of samples in the long term
18. Access to autoclave for sterilizing reusable equipment
Software and datasets
1. BioRender (https://www.biorender.com/). The following figures were created using BioRender: Graphical overview, Babler, K. (2024) https://www.BioRender.com/q34x703
Procedure
A. Wastewater collection
Wastewater can be collected as either a grab or composite sample. A discussion about the advantages and disadvantages of grab vs. composite samples in capturing the temporal and spatial variability is available in Babler et al. [11].
1. Prior to sample collection, obtain sterile collection bottles (minimum volume of 2 L for grab samples and 5 L for composite samples). The wastewater within these collection bottles will be split and transferred into containers so that a portion of the water can be analyzed in the field for basic water quality and another portion can be brought back to the laboratory for further processing (Graphical overview, Step 1).
Note 1: For “grab” sample type, the 2 L bottles should be fitted with a hose clamp attached to chains using two chain hooks to lower bottles down into sewer hole collection sites (Figure 1). During grab sample collection, lower the 2 L collection bottle into the sewer using its attached chains and fill with wastewater (Figure 1). Upon removal from the sewer, disinfect sample and chains by spraying with clean tap water followed by 99.5% isopropanol. Also, spray areas where wastewater may have spilled on the surface during sample collection.
Note 2: For “composite” sample type, the 5 L or larger bottles should be placed within an autosampler according to manufacturer’s instructions for appropriate sample collection by the autosampler. Following the completed run by the autosampler protocol, remove the bottle from the autosampler and immediately cap and disinfect the exterior collection bottle by first rinsing outside with clean water followed by spraying with 99.5% isopropanol.
2. Upon wastewater sample collection as either grab or composites, pour collected wastewater into two separate containers. Container 1: a sterile 500 mL plastic bottle with 0.5 mL of pre-dispensed sodium thiosulfate (use 100 g/L solution of sodium thiosulfate to provide 0.1 g/L) to reduce the chlorine residual. Container 2: a rinsed 500 mL plastic container deep enough for measurement of physical-chemical water quality parameters using the YSI probe. Label Container 1 and its lid and Container 2 with a paint pen prior to transferring the wastewater, as regular water-based markers or Sharpies will be removed by alcohol sprays used to disinfect the outside of the sample bottles after wastewater sample transfer. Upon transferring wastewater from the collection bottle into each container, the samples in the collection bottles are to be homogenized (shaken), if possible, and passed through a strainer and funnel to remove large solids (Figure 2). This pouring is to occur within a 5-gallon bucket to avoid spillage on the ground. Once pouring is finished, rinse the solids from the strainer and funnel with clean water and pour captured solids back into the sewer. Disinfect the strainer and funnel with isopropanol spray. Collection bottles can either be discarded or recycled. If recycled, they are to be capped and rinsed with clean water and then sprayed with isopropanol and placed in a separate leak-proof container for transport back to the laboratory for cleaning and sterilization.
3. For Container 1 (for laboratory processing), cap with lid, rinse the outside with clean water, then spray the outside with isopropanol. Place Container on ice or with frozen ice packs within a cooler for transport to the laboratory for further processing once collected.
4. For Container 2 (for field measurements of basic water quality parameters), take water quality measurements (e.g., pH, specific conductivity, dissolved oxygen, temperature; Figure 2B) on freshly collected wastewater sample in its own reservoir utilizing a pre-disinfected and pre-calibrated YSI Probe. Record collected data upon measurement. Discard the remaining wastewater after YSI measurement back into the sewer. Rinse the YSI probe with clean water into the bucket. Discard the rinse water into the sewer. Rinse the bucket with clean water and discard the rinse back into the sewer. Disinfect YSI and bucket with isopropanol spray.
Note: Water quality measurements are additional data that can be collected and are not necessarily a requirement to utilize this protocol. pH is the only necessary requirement (and can be completed in the laboratory) if intending to proceed to the RNA-specific filtration methods, which pretreats the sample with MgCl2 and HCl (lowering pH of the sample to a set range) (Section C). We do, however, recommend taking water quality parameters as environmental factors, such as water temperature, turbidity, pH, conductivity, or dissolved oxygen, that could impact pathogen recovery downstream. Therefore, having this data on a per-sample basis could be used to understand decreased pathogen recovery following molecular analysis.
Figure 1. Bottle chain configuration for grab sample collection. Configuration of sampling bottle with chain connection to facilitate lowering into the sanitary sewer for grab sample collection (A). Use of bottle and chain within a sanitary sewer (B).
Figure 2. Configuration of sample collection in the field. A strainer and funnel are combined to facilitate the removal of large solids while guiding the wastewater into a bottle (A). In the field, the funnel, strainer, and bottle are placed within a bucket to capture spillage of wastewater, avoiding contamination of the surrounding ground (B). In the right panel, a water quality sonde (YSI Probe) is placed within a bottle inside the bucket, which facilitates the collection of physical-chemical water quality data in the field.
5. Place laboratory samples (Container 1) within a BSL-2 hood upon arrival at the laboratory, homogenize by shaking, and level off to consistent volumes (500 mL) prior to pretreatment. (Note: 500 mL collection bottles have room for more volume in the wide-mouth part of the bottle; for grab samples especially, a volume >500 mL may be collected.) Pour any excess wastewater (i.e., volume >500 mL) into a sterile beaker to lower the water line to the level indicated on the pre-labeled bottle to the desired volume (500 mL). Discard the excess wastewater from the sterile beaker once the sample in the bottle is leveled. Autoclave beaker(s) after use.
Note: A pre-sterilized beaker is necessary in case excess sample is poured out, and some must be returned to the sample bottle to achieve a 500 mL volume. Use a different, sterile beaker per wastewater sample to avoid cross-contaminating samples.
Caution: For all steps within the laboratory while handling wastewater, appropriate PPE is to be worn (lab coat, gloves, disposable face mask, long pants, closed-toed shoes, and tied-back hair). Additional steps of disinfection with 99.5% isopropanol must be taken to ensure that no cross-contamination occurs between samples (i.e., work with one sample at a time, disinfect outside of sample bottle and equipment sample came in contact with, replace gloves, and disinfect any surface/equipment that was contaminated once you are finished), and sterile technique should be used for all processing steps. Before removing anything from the BSL-2 hood, be sure to disinfect using 99.5% isopropanol.
B. Preparation of biological recovery controls for addition to wastewater
Viral recovery control, OC43
1. For generating the OC43 viral control, Vero cells are grown in a T75 flask to 50% confluency and infected at an MOI 0.01 using a previously frozen aliquot of OC43 virus for 1.5 h in 12 mL of RPMI media at room temperature on a plate rocker.
2. After infection, add 12 mL of RPMI media to the flask and incubate cells at 37 °C overnight.
3. The following morning, remove media by aspiration and immediately replace with new media (12 mL). Incubate cells at 37 °C for four days.
4. Filter culture medium containing viral particles using a Pall 0.45-µm filter unit.
5. Purify total RNA from 50 μL aliquots of virus stock in quadruplicate and quantify by qPCR (absolute quantification alongside standards) to determine genomic copies per reaction of the viral stock.
6. Create aliquots that hold enough viral OC43 RNA to spike a desired amount of wastewater samples, per day of collection, at the desired concentration. Typically, the volume of recovery controls varies between 10 and 100 µL (Vvirus below). This volume is then multiplied by the number of samples expected per day and then by 1.2 to add another 20% volume to the viral aliquot volume to allow for adjustments on the day of sample collection. If the viral RNA spike is too concentrated, it can be diluted using RPMI media.
Note: To compute the volume of the spike for one sample, the concentration of the OC43 stock solution in genomic copies per microliter (Cvirus) is needed and provided through step B5. If, for example, the goal is to spike a volume of OC43, Vvirus, in microliters, to a set volume of wastewater, Vsample, in milliliters, to obtain 106 genomic copies per liter of wastewater sample, the equation to determine the spike volume is:
V
v
i
r
u
s
=
10
6
×
V
s
a
m
p
l
e
1000
×
C
v
i
r
u
s
The 1,000 in the above equation is used to convert from milliliters of wastewater sample to be spiked, Vsample, into units of liters.
7. Using a heat block, heat-inactivate OC43 viral aliquots at 56 °C for 15 min prior to adding to wastewater samples.
8. Add the OC43 recovery control to wastewater at a concentration of 106 genomic copies per liter (gc/L) of wastewater (Graphical overview, Step 1AB). Mix to homogenize. Following the closure of the bottle, spray bottle and surrounding area of the BSL-2 hood with 99.5% isopropanol to disinfect.
Cell recovery control, Mycobacterium smegmatis
9. Transfer Mycobacterium smegmatis from glycerol stock onto a LB agar plate containing 100 μg/mL ampicillin (for general selection and to ensure off-target bacteria do not grow, as M. smegmatis is resistant to ampicillin). Incubate at 37 °C for three days.
10. Inoculate a single, isolated M. smegmatis colony from the fresh plate into LB broth containing 0.05% Tween 80 and 100 μg/mL ampicillin and grow at 37 °C with shaking for 30 h.
11. Harvest cells at mid-log phase as determined by the optical density at 600 nm via analysis with a spectrophotometer (OD600 ~1.0 corresponds to a concentration of 1 × 108 cells per milliliter), and estimate the concentration of the M. smegmatis, CM.speg, in units of genomic copies per microliter (gc/μL).
12. Add M. smegmatis recovery control to wastewater at a concentration of 2 × 106 gc/L of wastewater (Graphical overview, Step 1AB). Mix to homogenize. Following the closure of the bottle, spray bottle and surrounding area of the BSL-2 hood with 99.5% isopropanol to disinfect.
Note: To compute the volume of the spike for one sample, the concentration of the M. smegmatis stock solution in genomic copies per microliter (CM.smeg) is needed and provided through step B11. If, for example, the goal is to spike a volume of M. smegmatis, VM.smeg, in microliters, to a set volume of wastewater, Vsample, in milliliters, to obtain 2 × 106 cells per liter of wastewater sample, the equation to determine the spike volume is:
V
M
.
s
m
e
g
=
2
×
10
6
×
V
s
a
m
p
l
e
1000
×
C
M
.
s
m
e
g
The 1,000 in the above equation is used to convert from milliliters of wastewater sample to be spiked, Vsample, into units of liters.
C. Additional pretreatment of wastewater for electronegative membrane filtration
Note: This step is recommended for viral RNA recovery.
1. Vortex the tube containing the biological controls (OC43 and M. smegmatis) to homogenize. Add the target volume of the biological control to the sample. Close the bottle and shake to homogenize.
2. Add magnesium chloride to a concentration of 50 mM and an autoclaved magnetic stir bar per sample bottle (Graphical overview, step 1B). If using 51% MgCl2, then add 9.4 mL per liter of eluate (= 4.7 mL for 500 mL wastewater sample). Close the bottle and shake to homogenize.
3. Place the wastewater sample bottle on a magnetic stir plate and insert the pre-calibrated (using pH standards 4, 7, and 10) pH probe to record the initial pH of the sample using a pH meter.
4. With the pH probe still inserted, use a sterile glass dropper to adjust the pH of wastewater sample to 3.5–4.5 by adding individual drops of hydrochloric acid (10%), being careful never to touch the dropper to the sample to avoid cross contamination. Record the number of drops used as well as the final resulting pH of the sample from the pH meter (Graphical overview, Step 1B).
5. Close the bottle and shake to homogenize.
D. Primary concentration by membrane filtration
1. Attach sterile Pall magnetic filter cup holders securely to the vacuum manifold (ensure that one Pall magnetic filter cup is used per sample to avoid cross-contamination between samples).
2. Remove the magnetic top cup. With a pair of sterile forceps (one that only comes in contact with sterile filter cups; usually kept within a small sterile beaker; autoclave after use), carefully place a MCE membrane filter (if electronegative filtration for both RNA and DNA recovery are chosen; Graphical overview, Step 2B) or GN-6 Metricel membrane filter (vacuum filtration without acid and magnesium chloride pre-treatment if recovering DNA only; Graphical overview; Step 2A) on the base of the filter cup (see General note 2). Reattach magnetic cup, turn on the vacuum, and open the vacuum manifold.
Note: MCE membrane filters are used for electronegative filtration, where samples have undergone a pH change and addition of salt (MgCl2) to help viral particles bind to the filter not only by size but also by electrical charge. Electronegatively charged MCE membrane filters are recommended for viral targets. GN-6 Metricel membrane filters are used for size-selection only and are recommended for use with bacterial and fungal targets for which DNA is extracted. Vacuum filtration with a GN-6 Metricel membrane filter can capture viruses if they adhere to particles larger than 0.45 µm in size. We recommend the preparation of two separate filters, one for RNA extraction (MCE) and another for DNA extraction (GN-6).
3. Use a sterile, sample-specific graduated cylinder (one per sample, autoclave after use) to measure incremental volumes of wastewater sample to pour into the Pall cup for filtration and concentration of the sample.
Caution: Depending on the total number of samples, a lot of equipment goes into successfully performing this procedure. Be sure to use pre-sterilized individual equipment per wastewater sample to avoid cross-contaminating sampling sites and ensure that sterile technique is employed for all sample processing. Pre-label all equipment and supplies that will be in direct contact with samples to avoid cross-contamination.
4. Add wastewater for filtration to the Pall cup (one per sample, autoclave after use) until the filter membrane is completely saturated and minimal flow through the membrane occurs. Record the total volume of wastewater filtered.
5. Remove the magnetic top cup and use a pair of sample-specific forceps (two per sample, autoclave after use) to handle the membrane (Figure 3). Sample-specific forceps should be held within 100 mL beakers to avoid cross-contamination with surfaces, other equipment, or different sample membranes (autoclave after use).
a. For DNA, a half-membrane is to be used for downstream analysis. To prepare the half-membrane, remove the membrane from the filter funnel base and place within a sterile Petri dish. Slice the filter in half with a sterile scalpel, then use sample-specific forceps to fold the membrane in on itself twice. Place half of the folded membrane directly into a 2 mL Zymo ZR BashingBead lysis tube (0.1 & 0.5 mm) containing 1 mL of 1× DNA/RNA Shield (pre-pipetted using sterile technique) and the other half in a second BashingBead lysis tube containing 1 mL of 1× DNA/RNA Shield OR in a 5 mL Eppendorf tube also containing 1 mL of 1× DNA/RNA shield to be used as backup or a technical replicate (Graphical overview, Step 2Ai–ii). The reason a half-membrane is used for downstream DNA analysis is due to the physical size limitations of the BashingBead lysis tubes.
b. For RNA, the whole-membrane is to be used for downstream analysis as the process is not limited by physical size limitations. Carefully fold the membrane in half and then in on itself two more times utilizing the same pair of sample-specific forceps. Place folded membrane directly into a 5 mL Eppendorf tube containing 2 mL of 1× DNA/RNA Shield (pre-pipetted using sterile technique) (Graphical overview, Step 2B).
Note: When performing RNA analysis workflows only, the volume of 1× DNA/RNA Shield can be reduced to 1.5 mL to increase concentration factors. We recommend the use of 2 mL of 1× DNA/RNA Shield to maintain the consistency of concentrates (1 filter per 2 mL of Shield) between the DNA workflow and the RNA workflow.
Figure 3. Filter concentration and placement within lysis buffer. After a recorded volume of wastewater is passed through for whole-membrane extraction, the filter is folded into itself three times (A) and then placed into a 5 mL centrifuge tube containing 2 mL of lysis buffer (Zymo DNA/RNA shield) (B).
6. Refrigerate wastewater filter concentrates at 4 °C (5 mL Eppendorf tube with full or half membrane or 2 mL Zymo ZR BashingBead lysis tube, 0.1 & 0.5 mm, with half membrane) until ready for further molecular processing. Of note, we do not recommend bead beating prior to RNA extraction as heat/friction may degrade viral nucleic acids plus bead beating releases ribosomal RNA from bacteria and higher microbes, which can interfere with viral RNA. We recommend chemical lysis provided from DNA/RNA Shield followed by slight vortexing for improved viral recoveries.
Pause point: The protocol can be paused following the completion of primary concentration, and the sample can be stored at 4 °C until ready for further analysis. We recommend conducting the extractions within 24 h.
7. Rinse and autoclave all equipment used to filter wastewater following its use for individual samples [e.g., Pall cups, forceps, beaker in which forceps were held, graduated cylinders, stir bars (if applicable)].
E. RNA extraction from wastewater concentrates
Note: This step is recommended for the extraction of RNA from viruses.
1. Prior to molecular analysis, ensure appropriate PPE is used and that benchtop and equipment is decontaminated with 70% ethanol.
2. Following a brief vortex to dislodge and slightly agitate collected solids captured by the membrane (Graphical overview, Step 3), briefly spin down to collect the liquid at the bottom of the tube and pipette 400 μL of wastewater concentrate from the 5 mL Eppendorf tube containing whole membrane into a new, sterile 1.7 mL microcentrifuge tube (Graphical overview, Step 4).
3. Use Zymo Quick-RNA Viral kit for RNA extraction. Add 800 μL of viral RNA buffer (2:1) to the 400 μL sample and mix well by repeated pipetting. All centrifugation steps are performed at 13,000× g.
4. Transfer the mixture, 600 μL at a time, over two separate steps, to a Zymo-Spin IC column in a new collection tube. Centrifuge for 2 min. Discard the flowthrough.
5. Add 660 μL of viral wash buffer to the column and centrifuge for 30 s. Discard the flowthrough. Repeat wash step.
6. Add 660 μL of 100% ethanol to the column and centrifuge for 1 min. Discard the flowthrough.
7. Elute the column into a sterile microcentrifuge tube with nuclease-free water.
Note 1: To compare head-to-head with DNA extraction from wastewater, elute with 100 μL. Continue to step E8 to perform PCR inhibitor removal.
Note 2: For standalone RNA analysis from wastewater, elute with 10 μL. Add 30 μL of HIV-1 RNA (approximately 100 copies/μL by qPCR) to the eluted sample to assess PCR inhibition. Alternative inhibition controls can be used as deemed appropriate. Additionally, combine 10 μL of nuclease-free water with 30 μL of HIV-1 RNA to use as a separate no-inhibition control sample. The sample is ready for qPCR analysis (skip steps E7–9).
8. Optional: Prepare a Zymo-Spin III-HRC column by inserting the column into a new collection tube. Add 600 μL of prep-solution from OneStep PCR Inhibitor Removal kit and centrifuge at 8,000× g for 3 min.
9. Place the prepared column into a new 1.7 mL microcentrifuge tube.
10. To remove qPCR inhibitors, transfer 100 μL of RNA eluate into Zymo-Spin III-HRC filters and centrifuge at 16,000× g for 3 min.
11. Store samples on ice for immediate analysis by qPCR.
Pause point: The protocol can be paused following the completion of RNA extraction, and samples can be stored at -80 °C until ready for further analysis.
F. DNA extraction from wastewater concentrates
1. Place 2 mL Zymo ZR BashingBead tubes with half membranes securely into an OMNI Bead-Ruptor 12 instrument and run protocol using the following parameters: 1 min bashing, 5 min rest (3× cycles, 18 min total run time) (Graphical overview, Step 3; see General note 3)
2. Centrifuge 2 mL Zymo ZR BashingBead tube at 12,000× g for 1 min to separate particles from solution following bead beating (Graphical overview, Step 4).
3. Transfer 400 μL of supernatant to a Zymo-Spin III-F filter from the ZymoBIOMICS DNA Miniprep kit placed in a clean collection tube and centrifuge at 8,000× g for 1 min. Discard the flowthrough.
4. Follow the remainder of ZymoBIOMICS DNA Miniprep kit protocol starting with the addition of 1,200 μL of ZymoBIOMICS DNA binding buffer.
Pause point: The protocol can be paused following the completion of the ZymoBIOMICS DNA Miniprep kit, and the sample can be stored at -20 °C until ready for further analysis.
G. Volcano 2nd Generation qPCR for quantifying RNA targets
1. Amplify purified RNA directly by qPCR using Volcano2G polymerase in 40 μL reactions with a 4 μL sample input (Graphical overview, Step 5). Prepare a master mix for the number of samples plus an additional 2 according to the following ratios:
a. Nuclease-free water (23.6 μL)
b. 1.1× Volcano buffer (8.8 μL of 5× stock)
c. 200 nM dNTPs (0.8 μL of 10 mM stock)
d. 2 units Volcano2G polymerase (0.4 μL of 5 units/μL)
e. 1 unit anti-Taq antibody (0.2 μL of 5 units/μL)
f. 500 nM target-specific forward primer (Table 1) (1.0 μL of 20 μM stock)
g. 500 nM target-specific reverse primer (Table 1) (1.0 μL of 20 μM stock)
h. 250 nM target-specific probe (FAM or HEX) (Table 1) (0.1 μL of 100 μM stock)
i. 1× Rox (0.1 μL of 400× stock)
Table 1. Optimized target-specific primer and probe sequences for V2G-qPCR and qPCR. Primer pairs (f and r) and target-specific probe sequences for all qPCR targets from previous publications utilizing this procedure and approach for analysis.
Molecular target Primer/probe Sequence of primer/probe 5' → 3'
SARS-CoV-2
CV3b/f
CV3c/r
CV3.prb
TGCTAACAAAGACGGCATCA
GTAGCACGATTGCAGCATTG
ACA TTG GCA CCC GCA ATC CTG CT (FAM)
Beta-2 Microglobulin
qB2M/f
qB2M/r
B2M.prb
CAAGGACTGGTCTTTCTATCTCTTGTAC
CTGCTTACATGTCTCGATCCC
CAAAGTCACATGGTTCACACGGCAG (FAM or HEX)
OC43
OC43/f
OC43/r
OC43.prb
CAACCAGGCTGATGTCAATAC
AAACCTAGTCGGAATAGCCTCA
ACATTGTCGATCGGGACCCAAGT (FAM or HEX)
HIV-1
RTwt3/f
V106/r
RT1.prb
GAAAATTAGTAGATTTCAGAGAACTTAATAAGAGAAC
CATCACCCACATCCAGTACTGTTA
TTCTGGGAAGTTCAATTAGGAATACCACATCCCGCAGG (FAM)
SIV LTR
SIV876/f
SIV999/r
SIV.prb
GCTAGACTCTCACCAGCACTTG
CTAGGAGAGATGGGAACACACA
TCCACGCTTGCTTGCTTAAAGACCTCT (FAM)
Mpox
Mpox/f
Mpox/r
Mpox.prb
TCTTGCTATCACATAATCTGRAAGCGTA
GATATAGCACCACATGCACCA
AAGCCGTAATCTATGTTGTCTATCGTGTCC (HEX)
PMMoV
PMMoV/f
PMMoV/r
PMMoV.prb
AGTGGTTTGACCTTAACGTTTGA
CCTACGTCTGACGACACAATCT
CCTACCGAAGCAAATGTCGCACT (HEX)
M. smegmatis
qMsmKat/f
qMsmKat/r
MsmKat.prb
CCGCTCGAAGAGGTCG
GTCCAGGTGACCTCGAGAC
TCCTTGCCGACGCCGGTG (HEX)
2. Briefly vortex and centrifuge master mix solution and pipette 36 μL of master mix into individual wells within a 96-well Bio-Rad hard shell plate.
3. For samples, carefully add 4 μL directly into the master mix within each well. For no template controls, add 4 μL of nuclease-free water into the master mix within each well. No-template controls are equivalent to negative controls and should be analyzed for targets to confirm the lack of amplification signal. For standards, add 4 μL of target-specific standards into the master mix spanning five wells on the plate (ranging from 101–105 copies/μL).
4. Seal the plate firmly with Microseal B, remove perforated edges, and spin down the plate in a plate spinner for 15 s to collect liquid at the bottom of the wells.
5. Power up the CFX Connect instrument and specify run parameters per molecular target being analyzed (Tables 2–4).
Table 2. Thermocycling conditions for the V2G-qPCR reaction (SARS-CoV-2 and pepper mild mottle virus targets).
R2 values reported as ≥0.96 and efficiency reported between 95% and 100%.
Step Temp. (°C) Duration No. of cycles
Initial denaturation 88 30 s 1
Denaturation 88 5 s 45
Annealing Ta = 60 20 s
Extension 72 15 s
Table 3. Thermocycling conditions for the V2G-qPCR reaction (human coronavirus-OC43, HIV, SIV, and Beta-2 microglobulin targets). R2 values reported as ≥0.96 and efficiency reported between 95% and 100%.
Step Temp. (°C) Duration No. of cycles
Initial denaturation 88 30 s 1
Denaturation 88 5 s 45
Annealing Ta = 60 15 s
Extension 72 15 s
Table 4. Thermocycling conditions for the qPCR reaction (Mpox). R2 values reported as ≥0.96 and efficiency reported between 95% and 100%.
Step Temp. (°C) Duration No. of cycles
Initial denaturation 95 1 min 1
Denaturation 95 10 s 45
Annealing Ta = 60 20 s
Extension 72 15 s
6. Transform raw qPCR values back to gc/L by correcting for qPCR input (4 μL), volume of 1× DNA/RNA shield, and volume of wastewater used to prepare filter concentrate (taking into consideration if a half or full membrane was used). See Data analysis below.
H. qPCR for quantifying DNA targets
1. Amplify purified DNA directly by qPCR using 2× TaqMan Fast Universal PCR master mix in 30 μL reactions with a 5 μL sample input (Graphical overview, Step 5).
2. Prepare a master mix for the number of samples plus an additional 2 according to the following ratios:
a. 1× Fast Mix (15 μL of a 2× mix)
b. 500 nM target-specific forward primer (0.75 μL of a 20 μM stock)
c. 500 nM target-specific reverse primers (0.75 μL of a 20 μM stock)
d. 250 nM target-specific probe (0.075 μL of a 100 μM stock)
e. Nuclease-free water (8.5 μL)
3. Briefly vortex and centrifuge master mix solution and pipette 25 μL master mix into individual wells within a 96-well Bio-Rad hard shell plate.
4. For samples, carefully add 5 μL directly into the master mix within each well. For no template controls, add 5 μL of nuclease-free water into the master mix within each well. No template controls are equivalent to negative controls and should be analyzed for targets to confirm the lack of amplification signal. For standards, add 5 μL of target-specific standards into the master mix spanning five wells on the plate (ranging from 101–105 copies/μL).
5. Seal plate firmly with Microseal B, remove perforated edges, and spin down the plate in a plate spinner for 30 s to collect liquid at the bottom of the wells.
6. Power up CFX Connect instrument and specify run parameters per molecular target being analyzed (Table 5).
Table 5. Thermocycling conditions for the qPCR reaction (Mycobacterium smegmatis)
Step Temp. (°C) Duration No. of cycles
Initial denaturation 95 2 min 1
Denaturation 95 10 s 40
Annealing/Extension Ta = 61 25 s
7. Transform raw qPCR values back to gc/L by correcting for qPCR input (5 μL), volume of 1× DNA/RNA shield, and volume of wastewater used to prepare filter concentrate (taking into consideration if a half or full membrane was used). See data analysis below.
Data analysis
To transform the raw qPCR values back to gc/L of raw wastewater, the computations are as follows.
We define the output from the qPCR reactions as genomic copies per reaction (GCPR). This is derived from a Cq value that is related, through a standard curve, to genomic copies per reaction. To convert GCPR to genomic copies per original volume of wastewater (GCWW) in units of genomic copies per liter, the following equation is used:
G
C
W
W
=
G
C
P
R
×
V
C
×
V
E
×
1000
V
F
×
V
A
×
V
R
where:
VF = Volume of wastewater passed through the filter in milliliters (e.g., 60 mL)
VC = Volume of concentrate in microliters (e.g., 2,000 μL or 1,500 μL volume of DNA/RNA Shield that the folded filter was placed)
VA = Volume of aliquot of the concentrate used for extraction in microliters (e.g., 400 μL of concentrate was used for extraction).
VE = Volume of concentrated nucleic acid after extraction in microliters. If additional inhibition controls are added to the VE (e.g., HIV RNA), the VE is the sum of the original volume of concentrated nucleic acid plus the volume of liquid added. (e.g., 10 μL of concentrated nucleic acid was produced after extraction and, to this volume, 30 μL of HIV was added for a total VE of 40 μL of extract).
VR = Reaction volume used for the PCR reaction in microliters (e.g., 2 μL or 4 μL).
Note: The 1,000 in the above equation is used to convert VF from milliliters to liters.
The example calculation utilizing the above numbers follows as:
G
C
W
W
=
G
C
P
R
×
1500
×
40
×
1000
60
×
400
×
4
To assess PCR inhibition, quantities of spiked HIV-1 RNA in samples are determined by V2G-qPCR and compared to quantities detected in the control in which HIV-1 RNA was spiked into clean water. A shift to a Cq value higher (±2) than that determined for the water control reveals the magnitude of inhibition that occurs during PCR amplification.
To compute percent recovery:
For OC43, assuming an equivalent of 106 gc/L was added to the original wastewater sample, then for each sample for which OC43 was added, compute GCWW as above. Percent recovery = (GCWW for OC43/106 gc/L) × 100%
For M. smegmatis, the percent recovery is computed in a similar fashion; but in this case, 2 × 106 gc/L of M. smegmatis is added to the sample as per the protocol described. As above, compute the GCWW for each sample for which M. smegmatis was added. Percent recovery = GCWW for M. smegmatis/(2 × 106 gc/L) × 100%.
Normalization of raw qPCR data:
As indicated within Table 1, normalization targets like PMMoV and Beta-2 microglobulin (B2M) have been previously assessed [19,20] utilizing the presented workflows. PMMoV is a virus present in wastewater in high concentrations due to its common presence in human fecal waste, and B2M is a human cellular housekeeping gene that upregulates during inflammation or infection. Our findings indicate that normalization may have benefits at smaller sewersheds but does not help significantly with larger sewersheds. The benefits of normalization will depend upon the variability in the proportion of the wastewater that is represented by human waste relative to the variability of the normalizing measurement itself. We recommend that each researcher evaluate whether normalization is beneficial and utilize a normalization factor applicable to the research to which this protocol is adapted. Additionally, we recommend evaluating several targets’ abilities to provide sound normalization benefits.
Note: When analyzing data from multiple measurements, statistical tests of normality are recommended to determine if data generated is normal, log-normal, or non-parametric. Statistical tests employed downstream should be appropriate for the intended analysis, specific to the performed experiment, and adjusted for the hypotheses being tested. We recommend analysis with and without outliers, and we also recommend plotting data to further assess outliers as a means of assessing whether statistical tests should include them.
Validation of protocol
This protocol has been used and validated in the following research article(s). The filter concentration steps have been validated against ultracentrifugation [22] and against magnetic bead separation [6]. The V2G-qPCR has been validated against traditional RT-qPCR [6,22]. Additional comparisons between different workflows with cross-laboratory comparisons are provided in Zhan et al. [19] and Zhan et al. [21]. The protocol has been validated from detection limits of 102 gc/L to values upward of 107 gc/L. The reproducibility of this proposed workflow was evaluated by Babler et al. [3]. For viral RNA (without bead beating), reproducibility defined by the coefficient of variation was found to be 25% based upon measurements of SARS-CoV-2 and 15% for measurements based upon OC43. For DNA (with bead beating), reproducibility was found to be 14% based upon measurements of M. smegmatis. Recoveries (including sample splitting, concentration, lysis, extraction, and qPCR) were 20%, on average based upon measurements of OC43.
• Babler et al. [3]. Expanding a Wastewater-Based Surveillance Methodology for DNA Isolation from a Workflow Optimized for SARS-CoV-2 RNA Quantification. Journal of Biomolecular Techniques. (Figure 1, Figure 2, Table 1).
This protocol in its entirety is based on the publication by Babler et al. 2023; each of the individual steps of electronegative filtration (ENF; filtration with MCE membrane filters), vacuum filtration (VF; using GN-6 Metricel membrane filters), bead beating samples (or excluding), and the separate DNA- and RNA-based extraction methods compare the approaches head-to-head in a detailed manner. Five liters total of wastewater from three sampling sites (one hospital and two from a regional wastewater treatment plant) was collected for this study, where four experimental conditions (ENF with and without bead beating and VF with and without bead beating) were assessed by qPCR downstream. The viral particle spiked-in control, analyzed downstream by qPCR, was human coronavirus-OC43, and the DNA spiked-in control was Mycobacterium smegmatis.Figure 1 shows the laboratory workflow performed for analysis, and Figure 2 illustrates the resulting quantification of three molecular targets: SARS-CoV-2, M. smegmatis, and OC43. Table 1 provides the specific primers and probes used by qPCR for detecting the molecular targets of focus. Shapiro-Wilk normality tests showed that the majority of the data was normally distributed, and no outliers were observed. As a result, paired t-tests and Pearson correlations were utilized to assess the differences among the methods used. Results of this study showed that the highest recoveries for SARS-CoV-2 corresponded to the VF filter concentrates excluding bead beating, followed closely by the ENF filter concentrates excluding bead beating. Statistical differences were observed between concentrates that underwent bead beating and those that did not with a significant loss in SARS-CoV-2 signal by qPCR with the inclusion of bead beating. For the spiked-in targets, OC43 was similar to SARS-CoV-2 in that bead beating resulted in a significant decrease in RNA detection by qPCR; VF non-bead beat samples resulted in the highest detection by qPCR with a significant difference between the concentrates beat vs. not beat. For the ENF concentrates, bead beat concentrates provided higher recoveries than non-bead beat concentrates. Future research was recommended to further evaluate the differences between the filtration methods. These results validate this protocol by providing replicable results and effective detection utilizing all steps of this listed procedure, including the mentioned controls.
• Babler et al. [14]. Detection of the clinically persistent, pathogenic yeast spp. Candida auris from hospital and municipal wastewater in Miami-Dade County, Florida. Science of the Total Environment (Figure 2, Table 1).
This publication utilized the vacuum filtration (coupled with GN-6 Metricel membrane filters) portion of this procedure and the following DNA extraction process utilizing the ZymoBIOMICS DNA Miniprep kit coupled with bead beating. Two sampling locations were of primary focus for this manuscript, a hospital housing C. auris patients and the corresponding regional wastewater treatment plant. Samples analyzed were part of a weekly surveillance program that spanned from May to September 2022 (38 total 500 mL samples, 19 total weeks from both sites). Routine SARS-CoV-2 data (and patient cases), as a part of the routine surveillance efforts, were also assessed alongside the C. auris target to determine if there was a trend between hospital cases of illness and positive signal at both wastewater sites. The C. auris target was analyzed by qPCR and compared against positive patient cases admitted to the hospital. No spiked-in controls were utilized for this publication; however, wastewater was separately filtered to culture C. auris from wastewater (not mentioned within this protocol), where grown colonies were isolated and analyzed by ClustalW sequence alignment after target amplification and Sanger Sequencing to confirm that they matched the C. auris genome described in GenBank at the NCBI. Pearson and Spearman correlation assessments were performed with the data set and compared against clinical case data for infected patients provided by the University’s hospital. Figure 2 illustrates a time series plot containing daily clinical data as well as wastewater signal (detected by qPCR) for C. auris as well as SARS-CoV-2. Table 1 provides the specific primers and probes used by qPCR for detecting the molecular targets of focus. Insignificant correlations were illustrated between sampling sites for given weeks and between clinical data and wastewater for these time periods. Future research was recommended based on the results, yet this publication provided valuable data in that C. auris was detectable from wastewater utilizing the above methodology and was viable enough for culture using additional methodologies (not described here).
• Sharkey et al. [17]. Monkeypox viral nucleic acids detected using both DNA and RNA extraction workflows. Science of the Total Environment. (Figure 2, Figure 3, Table 1).
The entirety of this protocol was also utilized for this publication, in which the electronegative filtration approach (MCE membrane filter not coupled with bead beating) and the VF approach utilizing bead beating were employed. The long-standing WBS program at the University of Miami employed the filtration approach with MCE membrane filters, and for this investigation developed a similar approach of vacuum filtration using GN-6 Metricel membrane filters for the express purpose of detecting DNA pathogens, such as Mpox. Weekly samples from the hospital as well as wastewater treatment plant were utilized spanning the outbreak period in Miami-Dade County of Mpox (June to September 2022). Spiked-in controls, OC43, and M. smegmatis were utilized to assess the validity of each workflow using ENF combined with the described RNA extraction and VF combined with the described DNA extraction. Clinical data was also utilized, since this investigation was a response to a public health outbreak with positive patient cases at the hospital samples for wastewater. Figure 2 illustrates the genomic copies per liter (gc/L) of Mpox DNA from wastewater detected by qPCR, and Figure 3 illustrates the gc/L of Mpox RNA, likely attributable to the detection of Mpox RNA being expressed in infected human cells that had been introduced into the wastewater. This study followed a detection-based approach. The correspondence focused on the presence of Mpox within wastewater and evaluated if that aligned with dates of Mpox-positive patients in the hospital. This study validates this listed procedure by providing evidence for the ability to use these methods to detect DNA viruses of concern such as Mpox.
• Babler et al. [11]. Degradation rates influence the ability of composite samples to represent 24-hourly means of SARS-CoV-2 and other microbiological target measures in wastewater. Science of the Total Environment (Figure 1, Figure 2, Figure 3).
This work used the MCE membrane filter and electronegative filtration process of this procedure, coupled with the RNA extraction portion, in a thorough investigation of timepoints to assess changes in microbial target concentrations. Three separate experiments, corresponding to a community scale (wastewater treatment plant, WWTP), cluster-of-buildings scale (connected by the same sewer line), and building scale sewershed sampling site, collecting 24 samples for assessment by V2G-qPCR were performed (n = 72 collections, n = 4 composites, three collected and one given by the wastewater treatment plant, “B” set of experiments). Additionally, at the start of each experiment, 16 L of wastewater was collected and split into 24 aliquots to be assessed individually alongside hourly collections from the sewer system (n = 72 aliquots from 3× large collections, “A” set of experiments). The procedure followed in this publication slightly modified the total volume of wastewater collected for necessary splits for assessing water quality, creating a composite from every hourly collected sample, and assessing every hourly sample with filtration. The filtration-based procedure, however, was not modified, and each hourly sample was able to be pretreated and filtered within the allotted time. The OC43 spiked-in control was utilized per sample, with an additional viral control of SIV also being added to assess degradation over time. Figure 1 illustrates sample splitting and analysis performed on the large collection (16 L) performed at the start of each experiment. Figure 2 illustrates sample splitting and analysis performed on the individual collections, taken every hour from the sewer system per sewer shed. As SARS-CoV-2 degradation and stability were the primary focus of this study, Figure 3 illustrates the gc/L of SARS-CoV-2 detected from each hourly sample, the aliquoted hourly assessed sample, and composites created per experiment for each scale of sample collection. Table 1 provides the specific primers and probes used by qPCR for detecting the molecular targets of focus. Shapiro-Wilk normality tests were performed on all qPCR datasets. Non-normal distributions were determined per molecular target assessed (all targets: SARS-CoV-2, SIV, PMMoV, B2M, HIV, and OC43); therefore, Spearman correlation tests were used to evaluate associations between variables over time. Degradation rates were determined per target and per experiment from the slope of the best-fit line between the natural logarithm of the target concentration at hour T divided by the initial target concentration. Moreover, one-sample t-tests were also performed to evaluate whether the mean of the hourly sample (“A” experiments) was statistically different than the created composite for that experiment. For the “B” experiments, one-sample Wilcoxon signed rank tests and one-sample t-tests were also performed to compare means between the grab samples and for the effective comparison of the 24 collected samples with the created composite. Additionally, the homogeneity of variance was evaluated using Levene’s tests for assessing the level of variability between each sewer shed scale from hour to hour. Ultimately, the stability of SARS-CoV-2, B2M, and PMMoV were illustrated by the assessment performed here over the course of 24 hours. SIV degraded significantly over the 24-hour timeframe, providing evidence that the other targets remained stable. Hourly variability between sewersheds and collected samples further supported that at any point in time, the levels of pathogens found within wastewater can be significantly higher or lower. These results solidify the recommendation of composite sampling to obtain a better estimate of target levels within any given day within a sewer system. The reproducibility of this study and thorough investigation into several different molecular targets validated the long-standing approach used by the University of Miami, described here, which employs pretreatment coupled with MCE membrane filter and electronegative filtration followed by RNA extraction and V2G-qPCR assessment.
• Babler et al. [6]. Comparison of Electronegative Filtration to Magnetic Bead-Based Concentration and V2G-qPCR to RT-qPCR for Quantifying Viral SARS-CoV-2 RNA from Wastewater. ACS ES&T Water (Figure 1, Figure 2, Figure 3, Table 2).
The purpose of this publication was to compare the MCE membrane filter coupled with electronegative filtration against another method of primary concentration, utilizing magnetic beads to determine the overall validity and effectiveness of employing filtration-based approaches for WBS. A total of 13 samples (corresponding to 5 total weeks and multiple study sites) were utilized for this experiment, and the RNA extraction process described above, followed by V2G-qPCR and a mainstream RT-qPCR approach, were employed. The additional comparison of V2G-qPCR against RT-qPCR was to validate the qPCR assay, since commercial kits (especially RT-qPCR kits) during the COVID-19 pandemic were difficult to obtain, keep stocked, and used consistently as many labs employed their use. Figure 1 visually illustrates the workflow performed in head-to-head comparison of the two primary concentration methods and qPCR analyses. Figure 2 provides the correlations between the electronegative filtration and magnetic bead-based approaches for SARS-CoV-2, B2M, and OC43, utilized here as a spiked-in control. Furthermore, Figure 3 provides the correlations for three SARS-CoV-2 genes (N3, N1, and ORF1ab) between the two qPCR approaches. Shapiro-Wilk normality assessments were run on each set of data per concentration method and molecular target. Spearman correlations were used to compare the log-transformed viral concentrations between the primary concentration and qPCR methods investigated. Mann-Whitney U tests were also used to evaluate whether the means for the data sets were statistically equivalent. Ultimately, the ability to detect SARS-CoV-2 and B2M was statistically equivalent between the two primary concentration methods, illustrating the use of both methodologies as sound strategies for detecting these molecular targets within wastewater. OC43 was generally better detected by electronegative filtration than by the magnetic bead-based approach, which further validates the use of filtration methodologies with the intended use of inactivated, spiked-in viruses. Between the qPCR approaches, correlations provided statistical equivalency for the V2G-qPCR approach with mainstream RT-qPCR approaches describing the experimental soundness of the assay. This publication validates the procedure listed here as a sound methodology, which is equivalently comparable to another commonly used primary concentration approach (magnetic-bead based). Moreover, this study validates the use of V2G-qPCR as an effective alternative to RT-qPCR for quantifying nucleic acids from wastewater.
General notes and troubleshooting
General notes
1. This protocol did not discuss the processing of DNA viruses. For DNA viruses, use steps in section E but with the ZymoBIOMICS DNA Miniprep kit instead of the Zymo Quick-RNA Viral kit.
2. In some cases, the addition of acid as part of electronegative filtration may contribute to the decay of nucleic acids. If recoveries are low for viruses, and wastewater solids are visible, consider omitting the HCl and MgCl2 addition, as there is a possibility that the viruses may have preferentially partitioned toward the solids and can therefore be removed without charge attraction.
3. If the HCl and MgCl2 addition is a concern for DNA recovery of bacteria and higher organisms, consider splitting the wastewater sample before the HCl and MgCl2 addition step. If DNA viruses are to be also analyzed, we recommend testing to determine whether the HCl and MgCl2 addition improves viral DNA recovery.
4. If recoveries are low for viruses, consider alternative concentration methods such as magnetic bead-based technologies or ultracentrifugation methods. These alternative concentration methods may be necessary for samples with very low suspended solids.
5. Biological controls recommended in this protocol to assess overall process recoveries are OC43 for RNA viruses and M. smegmatis for microbes with cell walls. Other viral, bacterial, fungal, and protozoa controls can be utilized instead and may be more applicable depending on the target nucleic acids.
6. MCE membrane filters are negatively charged. For electronegative filtration, salts (e.g., magnesium chloride) and acids (e.g., hydrochloric acid) are added to the sample to change the charge of viral particles to positive. These positively charged viral particles are then attracted to the negatively charged MCE membrane filters and thus captured by charge instead of by size. We have compared results both with electronegative filtration using MCE membrane filters along with regular vacuum filtration using GN-6 Metricel membrane filters (without salt and acid addition), and we obtained similar viral recoveries.
7. Both Pall MCE filters and GN-6 Metricel filters have the same pore size (0.45 µm) and both filters can also remove viruses by size exclusion if the they partition toward larger particulates that are common in wastewater. Therefore, GN-7. Metricel filters without acid or MgCl2 (as listed in 1 above) may be suitable for virus capture in wastewater. For viruses in dilute suspensions, evaluate the suitability of both filtration methods, as filtration processes even with charge capture (e.g., electronegative filtration), may not be suitable in their capture on a membrane.
8. Regarding the inclusion or exclusion of bead beating, researchers should consider the heat and friction naturally included with the process and determine if there is a positive or negative impact on the downstream yield or quantity of the sample/targets being analyzed. Bead beating or mechanical disruption of complex cell walls is necessary to release nucleic acids from bacteria, fungi, and protozoa. Bead beating may not be necessary for viruses as off-the-shelf lysis solutions such as DNA/RNA shield are capable of chemically lysing most viruses. Bead beating may be counter-productive to viral recoveries in heterogeneous mixtures such as wastewater. Bead beating releases nucleic acids from most microbes in wastewater including highly abundant ribosomal RNA, which may interfere with the ability to detect viral genetic targets [3]. As a result, we do not recommend bead beating if the focus is on analyzing viral targets. Additional experimentation may be necessary for specific matrix and target combinations.
Troubleshooting
Problem 1: Low flow in sewer hole, making desired volume of collection difficult.
Possible cause: In these cases, there are specialized sample collection systems, such as cutting off the screw top portion of the 2 L sample collection bottles to allow the bottle to lie flatter and closer to the bottom of the sewer pipe. Alternatively, there are sample collection nozzles that have been designed to capture water in as little as 1 cm of flow depth. These specialized nozzles are generally weighted and create a vacuum with inlets below the 1-cm threshold, allowing for peristaltic pumps of composite samplers to retrieve wastewater.
Solution: If the problem keeps occurring, consider changing the location and/or timing of sampling collection.
Problem 2: pH of wastewater sample difficult to lower with HCl.
Possible cause: Highly buffered sample, which may be impacted by turbidity and has been observed especially when sewers are undergoing construction due to suspected entrainment of calcium carbonate.
Solution: Adjust pH to range appropriately as possible but do not overload the sample with HCl for risk of degrading pathogens of interest. Proceed with a higher pH if necessary and document.
Problem 3: pH of wastewater sample already below the target pH.
Possible cause: Acid added to wastewater at the source.
Solution: Record the pH. Proceed with a lower pH and document.
Problem 4: Very turbid and particulate-filled wastewater sample.
Possible cause: Sampling sites like wastewater treatment plants can have unusually turbid and particulate-filled wastewater samples on any given day, dependent on the activities of the general public, weather patterns, etc.
Solution: When utilizing filtration-based approaches, if a sample is generally darker or more opaque-looking, it is a good rule of thumb to start with a low volume and then gradually increase until the membrane becomes clogged and saturated. Some samples/sampling sites can saturate a membrane with 5–10 mL when that site normally would saturate at 30–70 mL.
Problem 5: Very clear and translucent wastewater sample.
Possible cause: For sampling sites that do not receive much waste from households or that are close to bodies of water (i.e., lakes and oceans), wastewater samples can look no different from non-potable water.
Solution: Samples that are exceptionally clear and do not visually look like wastewater require higher volumes to saturate membranes. Volumes up to 150–200 mL need to be filtered to appropriately saturate a membrane.
Problem 6: Low recoveries of nucleic acid following bead beating of membranes.
Possible cause: Unoptimized membrane size where tube/beads cannot rupture membrane enough, or bashing protocol generating too much heat, or friction, which can degrade pathogens in wastewater.
Solution: Consider testing different cycling parameters for each specific bead-beating machine and perform multiple head-to-head comparisons of sample yield. If a membrane is too destroyed (e.g., powder-like), it can have a negative impact on yield, similarly to if the membrane is not shredded enough (e.g., large membrane particles/chunks) where particulate is not released from the membrane into solution.
Problem 7: Inhibition detected from the inhibition control.
Possible cause: The chemical make-up of the sample is interfering with the PCR process.
Solution: Consider running dilutions of the sample in an attempt to dilute out the inhibitors. Alternatively, consider integrating inhibitor clean-up (e.g., OneStep PCR Inhibitor Removal kit, as described in step E8). Also, adjusting the elution volume from nucleic acid extraction columns may also help to overcome inhibition (see Sharkey et al. [22] for details).
Problem 8: Negative values for target.
Possible cause: Could be a false negative or truly a negative.
Solution: Check if positive controls OC43 and M. smegmatis were recovered at anticipated levels. Also, check for the PMMoV and B2M normalization targets, which should be present in wastewater with human fecal inputs.
Acknowledgments
This work was supported financially by the National Institute On Drug Abuse of the National Institutes of Health (NIH) under Award Number U01DA053941 and by NIH P30A1073961 awarded to the University of Miami Center for AIDS Research. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health. Additional support for this study was received by the 4Catalyzer Foundation and through the University of Miami (Coral Gables, FL) administration. In-kind contributions were provided by University Facilities, University Environmental Health and Safety, and University of Miami Health Safety Division. Laboratory facilities and support were made available in-kind through the Sylvester Comprehensive Cancer Center, the Miami Center for AIDS Research, the Miami Clinical and Translational Science Institute, and the University of Miami College of Engineering.
This protocol was adapted and modified from Babler et al. [3] (doi: 10.7171/3fc1f5fe.dfa8d906).
Competing interests
The authors of this manuscript declare no competing interests.
Ethical considerations
This protocol does not involve human subjects nor the use of vertebrate animals.
References
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Article Information
Publication history
Received: Jul 5, 2024
Accepted: Dec 8, 2024
Available online: Jan 14, 2025
Copyright
© 2025 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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Environmental science
Microbiology > Pathogen detection
Molecular Biology
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5,190 | https://bio-protocol.org/en/bpdetail?id=5190&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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This is an In Press version of the protocol that has not yet been assigned to an issue.
Peer-reviewed
Transgene-free Genome Editing in Grapevine
EB Edoardo Bertini
ED Erica D’Incà
SZ Stefania Zattoni
SL Sara Lissandrini
LC Luca Cattaneo
CC Clarissa Ciffolillo
AA Alessandra Amato
MF Marianna Fasoli
SZ Sara Zenoni
In Press, Available online: Jan 09, 2025
DOI: 10.21769/BioProtoc.5190 Views: 98
Reviewed by: Clizia VillanoIgnacio Lescano Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Horticulture Research Oct 2022
Abstract
CRISPR/Cas9 genome editing technology has revolutionized plant breeding by offering precise and rapid modifications. Traditional breeding methods are often slow and imprecise, whereas CRISPR/Cas9 allows for targeted genetic improvements. Previously, direct delivery of Cas9-single guide RNA (sgRNA) ribonucleoprotein (RNP) complexes to grapevine (Vitis vinifera) protoplasts has been demonstrated, but successful regeneration of edited protoplasts into whole plants has not been achieved. Here, we describe an efficient protocol for obtaining transgene/DNA-free edited grapevine plants by transfecting protoplasts isolated from embryogenic callus and subsequently regenerating them. The regenerated edited plants were comparable in morphology and growth habit to wild-type controls. This protocol provides a highly efficient method for DNA-free genome editing in grapevine, addressing regulatory concerns and potentially facilitating the genetic improvement of grapevine and other woody crop plants.
Key features
• Protoplasts are one of the most commonly used systems for the application of new breeding technologies, including DNA-free genome editing.
• Protoplasts are a highly accessible platform by CRISPR-Cas9 ribonucleoparticles through chemical or physical transfection.
• CRISPR-Cas9 ribonucleoparticles avoid the use of both Agrobacterium tumefaciens and plasmids; no stable integration of exogenous DNA occurs.
• The genetic background of DNA-free edited plants regenerated from protoplasts remains unchanged and identical to the original plant.
Keywords: CRISPR/Cas9 Genome editing Grapevine Protoplasts DNA free Ribonucleoprotein complexes Embryogenic callus Plant regeneration
Graphical overview
Graphical overview of the workflow of DNA-free genome editing application in embryogenic calli-derived protoplasts and whole plant regeneration. Embryogenic calli are induced from leaves of in vitro plants or from stamens and pistils. After induction and proliferation, embryogenic calli are used for protoplast isolation. Pre-assembled CRISPR-Cas9 ribonucleoproteins (RNPs) are introduced into protoplasts by PEG-mediated transfection. Transfected protoplasts are cultivated using the disc-culture method. The regeneration of whole edited plants from transfected protoplasts occurs through somatic embryo formation.
Background
Grapevine (Vitis vinifera L.) is a significant global fruit crop, valued for its fresh produce and especially for wine. Genetic improvement of grapevine is crucial to address challenges such as climate change, higher demands for quality and quantity, and the need for product differentiation [1,2]. Breeding is a long, time-consuming process, often requiring several years to develop new varieties. Despite the quicker development potential of genetically modified (GM) crops, they encounter opposition from both the public and regulatory agencies due to concerns regarding health and environmental impacts [3]. Nowadays, new breeding technologies, particularly genome editing through the CRISPR/Cas9 system, provide a promising alternative by ensuring precise and targeted genetic modifications without altering the original genetic background. CRISPR/Cas9 uses the Cas9 endonuclease, guided to specific DNA sequences by a single guide RNA (sgRNA), to create site-specific double-stranded breaks. These breaks are typically repaired by non-homologous end joining, resulting in insertions or deletions, or by homology-directed repair if a donor template is available [4]. This method has been used to study and modify various genes in grapevine, enhancing traits such as disease resistance and plant architecture [5–9].
Most CRISPR/Cas9 applications in grapevine have involved stable integration of the editing machinery, thus creating GMOs. To comply with regulations requiring the absence of foreign DNA, two main strategies are used: removing CRISPR/Cas9 components after editing or directly delivering Cas9-sgRNA ribonucleoproteins (RNPs). The latter method avoids foreign DNA integration and has been demonstrated in grapevine protoplasts, but regenerating whole plants from these edited protoplasts has been challenging [10,11]. Recent advancements include both a procedure for regenerating whole plants from embryogenic callus-derived protoplasts in specific grapevine varieties [12] and a method for the regeneration of transgene-free edited grapevine plants from RNP-transfected protoplasts, confirming the feasibility of these approaches for producing normal and healthy transgene-free edited grapevine plants [13]. The advantages of this protocol include avoiding foreign DNA integration, thus addressing regulatory and public concerns associated with GMOs. Additionally, this method reduces the occurrence of genetic chimeras, which are common in organogenesis-based regeneration methods [14,15]. The approach is particularly suitable for varieties recalcitrant to traditional gene transfer methods, offering a versatile tool for grapevine breeding and functional genomics. Potential applications of this protocol extend beyond grapevine to other economically important woody and herbaceous crops. By refining the procedures for protoplast isolation, RNP transfection, and plant regeneration, this method could become a standard for developing transgene-free edited plants. This aligns with regulatory frameworks and facilitates broader acceptance of genome editing technologies in agriculture [16].
Materials and reagents
Biological materials
1. In vitro–grown grapevine plants
2. Inflorescences of open field plants or greenhouse-grown cuttings
Reagents
1. Enzymes
a. Cellulase R10 (Duchefa Biochemie, catalog number: C8001)
b. Macerozyme R10 (Duchefa Biochemie, catalog number: M8002)
c. Pectolyase Y-23 (Duchefa Biochemie, catalog number: P8004)
2. Fluorophores
a. Fluorescein diacetate (FDA) (Sigma-Aldrich, catalog number: 343209)
b. Fluorescent brightener 28 disodium salt (Chemcruz, catalog number: sc-218504)
3. Gelling agents
a. Agar TC (PhytoTech Labs, catalog number: A296)
b. Phytagel (Sigma-Aldrich, catalog number: P8169)
c. Gelrite (Duchefa Biochemie, catalog number: G1101)
4. Growth regulators
a. 6-Benzylaminopurine (6-BAP) (Duchefa Biochemie, catalog number: B0904)
b. 2,4-Dichlorophenoxyacetic acid (2,4-D) (Duchefa Biochemie, catalog number: D0911)
c. α-Naphthalene acetic acid (NAA) (Duchefa Biochemie, catalog number: N0903)
d. β-Naphthoxyacetic acid (NOA) (Duchefa Biochemie, catalog number: N0912)
5. Kit
a. QubitTM RNA BR Assay kit, 500 assays (Invitrogen, Thermo Fisher Scientific, catalog number: Q10211)
b. GeneArt Precision gRNA Synthesis kit (Invitrogen, Thermo Fisher Scientific, catalog number: A29377)
6. Proteins
a. TrueCut TM Cas9 protein v2 (Invitrogen, Thermo Fisher Scientific, catalog number: A36499)
7. Salts and biochemicals
a. CaCl2·2H2O (Duchefa Biochemie, catalog number: C0504)
b. Ca(NO3)2·4H2O (Duchefa Biochemie, catalog number: C0505)
c. KCl (Sigma-Aldrich, catalog number: 31219)
d. KH2PO4 (Duchefa Biochemie, catalog number: P0574)
e. KNO3 (Duchefa Biochemie, catalog number: P0519)
f. K2SO4 (Duchefa Biochemie, catalog number: P0535)
g. MgCl2·6H2O (Duchefa Biochemie, catalog number: M0533)
h. MgSO4·7H2O (Duchefa Biochemie, catalog number: M0513)
i. NaCl (Duchefa Biochemie, catalog number: S0520)
j. NH4NO3 (Duchefa Biochemie, catalog number: A0501)
k. NH4Cl (Duchefa Biochemie, catalog number: A0528)
l. CuSO4·5H2O (Duchefa Biochemie, catalog number: C0508)
m. FeNaEDTA (Duchefa Biochemie, catalog number: E0509)
n. FeSO4·7H2O (Duchefa Biochemie, catalog number: F0512)
o. H3BO3 (Duchefa Biochemie, catalog number: B0503)
p. MnSO·H2O (Duchefa Biochemie, catalog number: M0514)
q. Na2MO4·2H2O (Duchefa Biochemie, catalog number: S0525)
r. Na2EDTA·2H2O (Duchefa Biochemie, catalog number: E0511)
s. ZnSO4·7H2O (Duchefa Biochemie, catalog number: Z0526)
t. KI (Duchefa Biochemie, catalog number: P0518)
u. CoCl2·6H2O (Duchefa Biochemie, catalog number: C0507)
v. Sucrose (Duchefa Biochemie, catalog number: S0809)
w. D-Mannitol (Duchefa Biochemie, catalog number: M0803)
x. D-Glucose monohydrate (Duchefa Biochemie, catalog number: G0802)
y. Polyethylene glycol 4000 (PEG 4000) (Duchefa Biochemie, catalog number: P0804)
z. 2-(N-morpholino) ethane sulfonic acid (Duchefa Biochemie, catalog number: M1503)
aa. Biotin (Duchefa Biochemie, catalog number: B0603)
bb. Folic acid (Duchefa Biochemie, catalog number: F0608)
cc. Myo-inositol (Duchefa Biochemie, catalog number: I0609)
dd. Glycine (Duchefa Biochemie, catalog number: G0709)
ee. Nicotinic acid (Duchefa Biochemie, catalog number: N0611)
ff. Pyridoxine HCl (Duchefa Biochemie, catalog number: P0612)
gg. Thiamine HCl (Duchefa Biochemie, catalog number: T0614)
hh. D-pantothenate calcium (vitamin B5 calcium) (Duchefa Biochemie, catalog number: C0604)
ii. L-glutamic acid (Duchefa Biochemie, catalog number: G0707)
jj. L-phenylalanine (Duchefa Biochemie, catalog number: P0716)
kk. TopVision agarose (Thermo Scientific, Thermo Fisher Scientific, catalog number: R0491)
ll. 100 bp DNA ladder (Life Technologies, Thermo Fisher Scientific, catalog number: SM0241)
mm. Syber® safe DNA gel stain (Invitrogen, Thermo Fisher Scientific, catalog number: S33102)
nn. TriTrack DNA loading dye 6× (Life Technologies, Thermo Fisher Scientific, catalog number: R1161)
oo. Acetocarmine staining (Sigma-Aldrich, catalog number: 280370)
pp. NaClO (Sigma-Aldrich, catalog number: 1056142500)
qq. Tween-20 (Sigma-Aldrich, catalog number: P1379)
8. Supplements
a. Casein hydrolysate (Duchefa Biochemie, catalog number: C1301)
b. Activated charcoal (Sigma-Aldrich, catalog number: 31616)
Solutions
1. Macronutrients
a. NN macronutrients 10× (see Recipes)
b. MS macronutrients 10× (see Recipes)
c. C2D macronutrients 10× (see Recipes)
2. Micronutrients
a. NN micronutrients 100× (see Recipes)
b. MS micronutrients 1,000× (without FeEDTA) (see Recipes)
c. MS micronutrients 100× (see Recipes)
d. C2D micronutrients 1,000× (see Recipes)
3. Hormones
a. 2,4-Dichlorophenoxyacetic acid 1,000 µM (see Recipes)
b. 6-Benzylaminopurine 1,000 µM (see Recipes)
c. 1-Naphthaleneacetic acid 1,000 µM (see Recipes)
d. β-Naphthoxyacetic acid 1,000 µM (see Recipes)
4. Vitamins
a. NN vitamins 500× (see Recipes)
b. MS vitamins 500× (see Recipes)
c. C2D vitamins 1,000× (see Recipes)
d. B5 vitamins 500× (see Recipes)
e. Vitamins mix C1 500× (see Recipes)
f. Vitamins T 1,000×: (see Recipes)
5. Amino acids
a. Amino acids mix 1,000× (see Recipes)
6. Chemicals
a. FeEDTA 200× (see Recipes)
b. KCl 500 mM (see Recipes)
c. 2-(N-morpholino) ethanesulfonic acid 100 mM, pH 5.7 (see Recipes)
d. CaCl2·2H2O 1 M (see Recipes)
e. MgCl2·6H2O 500 mM (see Recipes)
f. NaCl 1 M (see Recipes)
7. Organics
a. Mannitol 1 M (see Recipes)
b. Glucose 3 M (see Recipes)
8. Culture media for induction of embryogenic calli (see Recipes)
a. NB2 (culture medium for induction of embryogenic calli from in vitro leaves)
b. PIV (culture medium for induction of embryogenic calli from stamens and pistils)
c. MSII (culture medium for induction of embryogenic calli from stamens and pistils)
9. Culture medium for long-term maintenance of embryogenic calli (C1P) (see Recipes)
10. Culture medium for somatic embryos full germination (see Recipes)
11. Culture media for somatic embryos shooting (see Recipes)
a. C2D
b. C2D4B
c. MG1
d. MG1–10B
12. Culture media for full plant development and rooting (see Recipes)
a. RIM
b. MSN
13. Culture media for protoplasts cultivation (see Recipes)
a. Solid culture medium
b. Liquid culture medium
14. Solution for protoplast isolation, purification, and transfection (see Recipes)
a. Digestion solution
b. Digestion solution without enzymes
c. Wash solution
d. W5 solution
e. MMG solution
f. PEG solution
Recipes
1. Macronutrients
a. NN macronutrients 10×: 2.2 g/L CaCl2·2H2O, 9.5 g/L KNO3, 7.2 g/L NH4NO3, 1.85 g/L MgSO4·7H2O, 0.68 g/L KH2PO4
b. MS macronutrients 10×: 16.5 g/L of NH4NO3, 4.4 g/L of CaCl2·2H2O, 3.7 g/L of MgSO4·7H2O, 19.7 g/L of KNO3 and 1.7 g/L of KH2РО4
c. C2D macronutrients 10×: 16.5 g/L NH4NO3, 19 g/L KNO3, 3.7 g/L MgSO4·7H2O, 1.7 g/L KH2PO4, 7.1 g/L Ca(NO3)2·4H2O, 0.28 g/L FeSO4·7H2O, 0.37 g/L Na2EDTA
2. Micronutrients
a. NN micronutrients 100×: 2.5 mg/L CuSO4·5H2O, 3.67 g/L FeNaEDTA, 1 g/L H3BO3, 1.9 g/L MnSO4·H2O, 25 mg/L Na2MoO4·2H2O, 1g/L ZnSO4·7H2O
b. MS micronutrients 1,000× (without FeEDTA): 6.2 g/L H3BO3, 16.9 g/L MnSO4·H2O, 8.6 g/L ZnSO4·7H2O, 0.83 g/L KI, 0.25 g/L Na2MoO4·2H2O, 25 mg/L CuSO4·5H2O, 25 mg/L CoCl2·6H2O
c. MS micronutrients 100×: 0.62 g/L H3BO3, 3.67 g/L FeNaEDTA, 1.69 g/L of MnSO4·H2O, 0.86 g/L ZnSO4·7H2O, 0.083 g/L of KI, 0.025 g/L Na2MoO4·2H2O, 0.0025 g/L CuSO4·5H2O, 0.0025 g/L CoCl2·6H2O
d. C2D micronutrients 1,000×: 0.64 g/L MnSO4·H2O, 6.2 g/L H3BO3, 8.6 g/L ZnSO4·7H2O, 0.25 g/L Na2MoO4·2H2O, 25 mg/L CuSO4·5H2O, 25 mg/L CoCl2·6H2O
3. Hormones
Note: All the following hormone powders are soluble in NaOH 1 M. It is important to add NaOH 1 M to the weighed powder before the addition of water.
a. 2,4-Dichlorophenoxyacetic acid 1,000 µM: 0,221 g/L of 2.4-dichlorophenoxyacetic acid
b. 6-Benzylaminopurine 1,000 µM: 0.2252 g/L of 6-benzylaminopurine
c. 1-Naphthaleneacetic acid 1,000 µM: 0.1862 g/L of 1-naphthaleneacetic acid
d. β-Naphthoxyacetic acid 1,000 µM: 0.2022 g/L of β-naphthoxyacetic acid
4.Vitamins
a. NN vitamins 500×: 25 mg/L biotin, 0.25 g/L folic acid, 1 g/L glycine, 50 g/L myo-Inositol, 2.5 g/L nicotinic acid, 0.25 g/L Pyridoxine HCl, 0.25 g/L thiamine HCl
b. MS vitamins 500×: 1 g/L glycine, 50 g/L myo-inositol, 0.25 g/L nicotinic acid, 0.25 g/L pyridoxine HCl, 0.05 g/L thiamine HCl
c. C2D vitamins 1,000×: 1 g/L thiamine HCl, 10 g/L myo-inositol, 1 g/L nicotinic acid, 1 g/L pyridoxine HCl
d. B5 vitamins 500×: 50 g/L myo-inositol, 5 g/L thiamine HCl, 0.5 g/L nicotinic acid, 0.5 g/L pyridoxine HCl
e. Vitamins mix C1 500×: 50 g/L of myo-inositol, 5 g/L of nicotinic acid, 5 g/L of thiamine-HCI, 0.5 g/L of pyridoxine-HCl, 0.5 g/L of calcium pantothenate, 0.005 g/L of biotin
f. Vitamins T 1,000×: 50 g/L myo-inositol, 1 g/L nicotinic acid, 1 g/L thiamine HCl, 1 g/L pyridoxine HC, 1 g/L calcium pantothenate, 0.01 g/L biotin.
5. Amino acids
a. Amino acids mix 1,000×: 100 g/L L-glutamic acid (monosodium salt), 10 g/L L-phenylalanine, 2 g/L glycine
6. Chemicals
a. FeEDTA 200×: 7.44 g/L of Na2EDTA·2H2O, 1.86 g/L of FeSO4·7H2O
b. KCl 500 mM: 37.775 g/L of KCL
c. 2-(N-morpholino) ethanesulfonic acid 100 mM, pH 5.7: 19.52 g/L of 2-(N-morpholino) ethanesulfonic acid.
Note: Adjust the pH of this solution to 5.7 with KOH 1 M
d. CaCl2·2H2O 1 M: 146.9 g/L of CaCl2·2H2O
e. MgCl2·6H2O 500 mM: 101.655 g/L of MgCl2·6H2O
f. NaCl 1M: 58.44 g/L of NaCl
7. Organics
a. Mannitol 1 M: 182.17 g/L of D-mannitol
b. Glucose 3 M: 594.6 g/L of glucose
8. Culture media for induction of embryogenic calli
a. NB2 (culture medium for induction of embryogenic calli from in vitro leaves)
Component Stock concentration Final concentration Quantity or Volume (1 L)
NN macronutrients 10× 1× 100 mL
NN micronutrients 100× 1× 10 mL
MS vitamins 500× 1× 2 mL
6-benzylaminopurine 1,000 μM 1.0 μM 1 mL
2,4-dichlorophenoxyacetic acid 1,000 μM 5.0 μM 5 mL
Myo-inositol / 0.1 g/L 0.1 g
Sucrose / 20 g/L 20 g
Adjust to final pH 6.0 with KOH 1 M
Agar TC / 7 g/L 7 g
b. PIV (culture medium for induction of embryogenic calli from stamens and pistils)
Component Stock concentration Final concentration Quantity or Volume (1 L)
NN macronutrients 10× 1× 100 mL
MS micronutrients 1,000× 1× 1 mL
FeEDTA 200× 1× 5 mL
B5 vitamins 500× 1× 2 mL
6-benzylaminopurine 1,000 μM 8.9 μM 8.9 mL
2,4-dichlorophenoxyacetic acid 1,000 μM 4.5 μM 4.5 mL
Sucrose 60 g/L 60 g
Adjust to final pH 5.7 with KOH 1M
Phytagel / 3 g/L 3 g
c. MSII (culture medium for induction of embryogenic calli from stamens and pistils)
Component Stock concentration Final concentration Quantity or Volume (1 L)
MS macronutrients 10× 1× 100 mL
MS micronutrients 100× 1× 10 mL
MS vitamins 500× 1× 2 mL
β-naphthoxyacetic acid 1,000 μM 2.5 μM 2.5 mL
6-benzylaminopurine 1,000 μM 5 μM 5 mL
2,4-dichlorophenoxyacetic acid 1,000 μM 2.5 μM 2.5 mL
Myo-inositol / 0.1 g/L 0.1 g
Sucrose / 20 g/L 20 g
Adjust to final pH 6.0 with KOH 1M
Agar TC / 7 g/L 7 g
9. Culture medium for long-term maintenance of embryogenic calli (C1P)
Component Stock concentration Final concentration Quantity or Volume (1 L)
MS macronutrients 10× 1× 100 mL
MS micronutrients 1,000× 1× 1 mL
Vitamins C1 500× 1× 2 mL
AA mix 1,000× 1× 1 mL
Fe-EDTA 200× 1× 5 mL
2,4-dichlorophenoxyacetic acid 1,000 μM 5 μM 5 mL
6-benzylaminopurine 1,000 μM 1 μM 1 mL
Casein hydrolysate / 1 g/L 1 g
Sucrose / 30 g/L 30 g
Adjust to final pH 5.8 with KOH 1 M
Phytagel / 5g/L 5 g
10. Culture medium for somatic embryos full germination
Component Stock concentration Final concentration Quantity or Volume (1 L)
NN macronutrients 10× 1× 100 mL
NN micronutrients 100× 1× 10 mL
NN vitamins 500× 1× 2 mL
Sucrose / 30 g/L 30 g
Adjust to final pH 5.8 with KOH 1 M
Gelrite / 2 g/L 2 g
11. Culture media for somatic embryo shooting
a. C2D
Component Stock concentration Final concentration Quantity or Volume (1 L)
C2D macronutrients 10× 1× 100 mL
C2D micronutrients 1,000× 1× 1 mL
C2D vitamins 1,000× 1× 1 mL
Sucrose / 30 g/L 30 g
Adjust to final pH 5.8 with KOH 1 M
Agar TC / 7 g/L 7 g
b. C2D4B
Component Stock concentration Final concentration Quantity or Volume (1 L)
C2D macronutrients 10× 1× 100 mL
C2D micronutrients 1,000× 1× 1 mL
C2D vitamins 1,000× 1× 1 mL
6-benzylaminopurine 1,000 μM 4 μM 4 mL
Sucrose / 30 g/L 30 g
Adjust to final pH 5.8 with KOH 1 M
Agar TC / 7 g/L 7 g
c. MG1
Component Stock concentration Final concentration Quantity or Volume (1 L)
NN macronutrients 10× 1× 100 mL
MS micronutrients 1,000× 1× 1 mL
Fe-EDTA 200× 1× 5 mL
B5 vitamins 500× 1× 2 mL
Sucrose / 30 g/L 30 g
Adjust to final pH 5.8 with KOH 1 M
Agar TC / 7 g/L 7 g
Activated charcoal / 2.5 g/L 2.5 g
d. MG1-10B
Component Stock concentration Final concentration Quantity or Volume (1 L)
NN macronutrients 10× 1× 100 mL
MS micronutrients 1,000× 1× 1 mL
Fe-EDTA 200× 1× 5 mL
B5 vitamins 500× 1× 2 mL
6-benzylaminopurine 1,000 μM 10 μM 10 mL
Sucrose / 30 g/L 30 g
Adjust to final pH = 5.8 with KOH 1 M
Agar TC / 7 g/L 7 g
Activated charcoal / 2.5 g/L 2.5 g
12. Culture media for full plant development and rooting
a. RIM
Component Stock concentration Final concentration Quantity or Volume (1 L)
MS macronutrients 10× 1× 100 mL
MS micronutrients 1,000× 1× 1 mL
Fe-EDTA 200× 1× 5 mL
Vitamins T 1,000× 1× 1 mL
1-naphthaleneacetic acid 1,000 μM 0.5 μM 500 μL
Sucrose / 30 g/L 30 g
Adjust to final pH 6.0 with KOH 1 M
Agar TC / 7 g/L 7 g
b. MSN
Component Stock concentration Final concentration Quantity or Volume (1 L)
MS macronutrients 10× 1× 100 mL
MS micronutrients 100× 1× 10 mL
MS vitamins 500× 1× 2 mL
1-naphthaleneacetic acid 1,000 μM 0.5 μM 500 μL
Sucrose / 30 g/L 30 g
Adjust to final pH 5.8 with KOH 1M
Agar TC / 7 g/L 7 g
13. Culture media for protoplast cultivation
a. Solid culture medium
Component Stock concentration Final concentration Quantity or Volume (50 mL)
NN macronutrients 10× 1× 5 mL
NN micronutrients 100× 1× 500 μL
NN vitamins 500× 1× 100 μL
1-naphthaleneacetic acid 1,000 μM 10 μM 500 μL
6-benzylaminopurine 1,000 μM 2 μM 100 μL
Glucose 3 M 0.3 M 5 mL
Sucrose / 30 g/L 1.50 g
Adjust to final pH 5.7 with KOH 1 M
Gelrite / 2 g/L 0.1 g
b. Liquid culture medium
Component Stock concentration Final concentration Quantity or Volume (50 mL)
NN macronutrients 10× 1× 5 mL
NN micronutrients 100× 1× 500 μL
NN vitamins 500× 1× 100 μL
1-Naphthaleneacetic acid 1,000 μM 10 μM 500 μL
6-benzylaminopurine 1,000 μM 2 μM 100 μL
Glucose 3 M 0.3 M 5 mL
Sucrose / 30 g/L 1.50 g
Adjust to final pH 5.7 with KOH 1 M
Activated charcoal / 3 g/L 0.15 g
14. Solution for protoplast isolation, purification, and transfection
a. Digestion solution
Component Stock concentration Final concentration Quantity or Volume (20 mL)
Cellulase R10 / 2% w/v 0.4g
Macerozyme R10 / 1 % w/v 0.2 g
Pectolyase Y-23 / 0.05 % w/v 0.01 g
CaCl2·2H2O 1 M 10 mM 200 μL
2-(N-morpholino) ethanesulfonic acid, pH 5.7 100 mM 5 mM 1 mL
Mannitol 1 M 0.5 M 10 mL
Adjust to final pH 5.7 with KOH 1 M
b. Digestion solution without enzymes
Component Stock concentration Final concentration Quantity or Volume (20 mL)
CaCl2·2H2O 1 M 10 mM 200 μL
2-(N-morpholino) ethanesulfonic acid, pH 5.7 100 mM 5 mM 1 mL
Mannitol 1 M 0.5 M 10 mL
Adjust to final pH 5.7 with KOH 1 M
c. Wash solution
Component Stock concentration Final concentration Quantity or Volume (50 mL)
CaCl2·2H2O 1 M 10 mM 500 μL
Mannitol 1 M 0.5 M 25 mL
Adjust to final pH 5.7 with KOH 1 M
d. W5 solution
Component Stock concentration Final concentration Quantity or Volume (40 mL)
CaCl2·2H2O 1 M 125 mM 5 mL
2-(N-morpholino) ethanesulfonic acid, pH 5.7 100 mM 2 mM 800 μL
NaCl 1 M 154 mM 6.16 mL
KCl 500 mM 5 mM 400 μL
e. MMG solution
Component Stock concentration Final concentration Quantity or Volume (10 mL)
Mannitol 1 M 0.4 M 4 mL
MgCl2·6H2O 500 mM 15 mM 300 μL
2-(N-morpholino) ethanesulfonic acid, pH 5.7 100 mM 4 mM 400 μL
f. PEG solution
Component Stock concentration Final concentration
Quantity or Volume
(10 mL)
Mannitol 1 M 0.2 M 2 mL
CaCl2·2H2O 500 mM 100 mM 1 mL
PEG 4000 / 40% 4 g
Laboratory supplies
1. Steri vent container 107 × 94 × 96 mm (Duchefa Biochemie, catalog number: S1682)
2. Nylon filter 60 μM (AGRINOVA)
3. Petri dishes 92 × 16 (VWR, catalog number: 391-0493)
4. Petri dishes 60 × 15 (Greiner, catalog number: 628161)
5. Petri dishes 35 × 15 (Thermo Scientific, catalog number: 153066)
6. Sterile needles 0.7 × 50 mm (Henke Sass Wolf, catalog number: 471,0007050)
7. Sterile syringes 1 mL (Terumo, catalog number: MDSS01SE)
8. Sterile syringes 50 mL (NIPRO, catalog number: BSS131)
9. Sterile syringes 20 mL (BD Plastipak, catalog number: 300613)
10. Sterile centrifuge tubes, Falcon 15 mL (VWR, catalog number: VWRI525-0607)
11. Sterile centrifuge tubes, Falcon 50 mL (VWR, catalog number: VWRI525-0612)
12. Pipettes (Eppendorf)
13. Fast-Read102® (Kova International, catalog number: BVS100H)
14. Filter 0.2 μm (Sarstedt, catalog number: 83.1826.001)
15. Pasteur 3 mL (Sarstedt, catalog number: D-51588)
16. Filter tips (2,5 μL, 20 μL, 200 μL, 1,000 μL, 5 mL)
17. Scalpel blades 21 and 11 (Duchefa)
18. Microscope slides
19. Coverslips
20. PCR tubes
21. 1.5 mL Eppendorf tubes
22. Sterile scalpels
23. Sterile forceps
24. Parafilm
Equipment
1. Heating magnetic stirrer (VWR®, model: 442-0664)
2. pH meter (CRISON, model: BASIC 20+)
3. Weighing balance (OHAUS, model: AX422/E)
4. Analytical balance (OHAUS, model: PA114C)
5. Glass microsphere sterilizer (AgnTho’s AB, model: steri 250, Art-Nr 31’101)
6. ChemiDoc imaging system (Bio-Rad, model: 12003153)
7. Thermal cycler (Bio-Rad, model: S1,000)
8. Horizontal laminar flow cabinet (BIOAIR, model: aura HZ72)
9. PowerPacTM basic power supply (Bio-Rad, model: 1645050)
10. Horizontal electrophoresis cells (Bio-Rad, model: Sub-Cell GT)
11. Horizontal electrophoresis cells (ELETTROFOR, models: OA-50; OA-78)
12. INCU-Line 150R (VWR®, model: 390-1338)
13. Benchtop microcentrifuge (Eppendorf, catalog number: 5420)
14. Centrifuge (Eppendorf, model: 5804 R)
15. Stereomicroscopes (Leica, models: MZ16 F; EZ4)
16. Optical microscope (Leica, model: DM2500)
Procedure
All the following steps must be done under a laminar flow hood in order to maintain sterile conditions unless otherwise specified. Be sure to use sterile laboratory supplies and to have an available bead sterilizer in order to be able to periodically sterilize all the tools used to handle the plant material.
Embryogenic calli induction and long-term maintenance
A. Embryogenic calli induction from unopened leaves
1. Open the container in which the in vitro grapevine plantlets are present (Figure 1A).
2. Cut the unopened leaves near the apex of the shoots with a scalpel.
3. Collect all the leaves inside a Petri dish using forceps.
4. Prepare some Petri dishes with NB2 solid medium inside.
5. Still using forceps, place up to five unopened leaves inside each NB2 Petri dish (Figure 1B), with one side fully adjacent to the induction medium (abaxial side).
6. Incubate the NB2 petri dishes prepared with the plant material in full darkness at 27 °C for 6–7 weeks.
7. Check weekly for the evolution of the plant material in NB2 medium and the induction of embryogenic calli (Figure 1L).
Figure 1. Embryogenic calli induction and proliferation. A. In vitro grapevine plantlet. The arrow indicates the apex. B. In vitro leaf in NB2 medium (scale bar: 6 mm). C. Inflorescence from greenhouse-grown cutting. D. Grapevine flower (scale bar: 3 mm). Grey arrow indicates the calyptra, and blue arrow indicates the calyx. E. Stamen (scale bar: 0.5 mm). F. Pistil (scale bar: 0.5 mm). Pollen microsporogenesis stages: mother cells (G), tetrads (H), uninucleate pollen cells (I) (scale bars: 50 μm). L. In vitro leaf-induced embryogenic callus in NB2 medium (scale bar: 2 mm). M: Stamen/pistil-induced embryogenic callus in MSII/PIV media (scale bar: 2 mm). N: Embryogenic calli proliferated in C1P medium (scale bar: 3 mm).
B. Embryogenic calli induction from stamens and pistils
1. Examination of the microsporogenesis development stage in anthers
Steps B1a–B1j can be performed in non-sterile conditions.
a. Collect a single flower from an inflorescence (Figure 1C).
b. Using a stereomicroscope, open the flower (Figure 1D) by cutting just below the calyptra with the sharp tip of a needle connected with a syringe in order to eliminate the calyx from the flower and maintain just the upper part with the calyptra, stamens, and pistil.
c. Still using a stereomicroscope, with two syringes, open the calyptra and reveal the stamens (Figure 1E) and the pistil (Figure 2F) inside.
d. Pick up all the stamens and place them on a microscope slide.
e. Crush the anthers in order to reveal the reproductive cells hidden inside (Figure 1G, H, I).
f. Soak the crushed material with 2–3 droplets of acetocarmine.
g. Cover the sample with a coverslip.
h. Observe the structures revealed in the crushed material under an optical microscope.
i. Repeat steps B1a–B1i by sampling different flowers from different parts of the inflorescence.
j. If all the observations show the microsporogenesis stage (Figure 1G, H, I) required, it is possible to proceed with the inflorescence sampling.
2. Inflorescence sampling and sterilization
a. Prepare the solution for the inflorescence sterilization (NaClO 3% and Tween-20 0.1%) in a Falcon tube.
b. Cut the entire inflorescence and immerse it in the sterilization solution.
c. Mix continuously by inversion for 10 min.
d. Discard the sterilization solution inside a collection container.
e. Fill the empty Falcon, which contains the inflorescence, with sterile double-distilled H2O.
f. Mix continuously by inversion for 5 min.
g. Discard the water inside the collection container.
h. Repeat steps B2e, B2f and B2g two more times.
i. Using forceps, gently pick the inflorescence from the Falcon and place it in a new Petri dish.
j. Store the sterilized inflorescence at 4 °C up to 72 h.
3. Flower explants cultivation
a. Prepare a Petri dish with single drops of sterile double-distilled H2O on different sides of the Petri dish.
b. Cut some flowers from the inflorescence with forceps and a scalpel and transfer them to the Petri dish prepared in the previous step.
c. Be sure to place the flowers inside sterile double-distilled H2O droplets.
d. Using a stereomicroscope, with the sharp tip of a couple of sterile needles connected to two syringes, apply a cut in the flower just below the calyptra and discard the lower part of the organ (calyx).
e. Using needles, eliminate the calyptra and reveal the stamens and the pistil.
f. Gently pick the stamens and the pistil with the smooth part of the tip of a sterile needle, and one by one place them in a Petri dish with solid induction media (MSII or PIV) inside.
g. Place 52 explants (16 pistils and 36 stamens) in each induction media Petri dish.
h. Prepare up to 10 plates for each embryogenic calli induction medium.
i. Incubate for 2–4 months in darkness at 27 °C.
j. Using the stereomicroscope, check weekly the formation of embryogenic calli (Figure 1M).
C. Embryogenic calli proliferation
1. Selection of embryogenic calli and transfer in C1P medium
a. Using a stereomicroscope, select white and granular shaped embryogenic calli (Figure 1M).
b. Mark the top of the Petri dish in correspondence with the explant selected for the propagation of the embryogenic calli.
c. Using a stereomicroscope, for each embryogenic callus selected (step C1b), collect all the embryogenic material emitted by the explant selected and transfer it to a new C1P Petri dish.
d. Prepare one or more C1P Petri dishes, each containing new embryogenic calli.
e. Using a scalpel, put embryogenic calli close to each other.
f. Repeat from step C1c to step C1e for each embryogenic callus (step C1b) selected.
g. Incubate the C1P petri dishes in darkness at 27 °C for 4 weeks.
2. First subculture in C1P medium and subsequent subcultures
a. Using a stereomicroscope, collect embryogenic material using a scalpel from 4-week-old C1P Petri dishes (Figure 2N).
b. Transfer selected embryogenic material to a new C1P Petri dish.
c. Prepare up to six new C1P Petri dishes.
d. Adjust the position of the embryogenic callus as described in step C1e.
e. Repeat from step C2a to step C2d for all embryogenic callus selected (step C1b).
f. Incubate in darkness at 27 °C for 4 weeks.
g. Repeat the subculture process every 4 weeks and prepare in each cycle six C1P Petri dishes with fresh embryogenic material for each embryogenic callus selected.
Protoplast isolation and cultivation
A. Preparation of embryogenic calli
1. Using a stereomicroscope, select compact and white cream-colored embryogenic callus from 4-week-old embryogenic culture masses grown in C1P medium.
2. Transfer them to fresh C1P medium.
3. Prepare up to seven C1P plates.
4. Incubate them at 27 °C in darkness for 7–10 days.
B. Protoplasts isolation
1. Using forceps, transfer embryogenic calli (Figure 2A) to the 60 × 10 mm Petri dish. Additionally, transfer a small amount of embryogenic calli as control to the 35 × 10 mm Petri dish.
2. Weigh embryogenic calli collected in the 60 × 10 mm Petri dish and add 10 g/mL of filtered-sterilized digestion solution.
3. Add 1 mL of filtered-sterilized digestion solution without enzymes in the 35 × 10 mm Petri dish.
4. Incubate both Petri dishes in darkness for 5–6 h on a rotating shaker.
Figure 2. Protoplast isolation from embryogenic calli, analysis of cell wall digestion, and protoplast viability. A. Embryogenic callus (7–10 days) in C1 medium (scale bar: 2 mm). B. Protoplasts isolated from embryogenic calli following 5 h of incubation with digestion solution. C, D. Protoplasts stained with fluorescent brightener 28 under white (C) and UV (D) light. The absence of blue signal indicates the correct cell wall digestion. E, F. Embryogenic calli (control) stained with fluorescent brightener 28 under white (E) and UV (F) light. The blue signal indicates the presence of the cell wall. G, H. Protoplasts stained with FDA under white (G) and UV (H) light. The green fluorescent signal indicates that protoplasts are viable and therefore can be used for transfection and cultivation (Scale bars: 20 μm).
C. Monitoring of progression of cell wall digestion
1. After 90 min, take an aliquot from the Petri dish containing calli in digestion solution (step B4) and observe the progress of isolation under an optical microscope.
2. Repeat step C1 after 3 h of incubation.
3. After about 5–6 h of incubation (Figure 2B), evaluate the correct digestion of the cell wall by staining both protoplasts and control with 2 μM fluorescent brightener 28.
4. Mix by pipetting.
5. Prepare a microscope slide and observe both protoplasts (Figure 2C) and control (Figure 2E) under UV light (Figure 2D, F).
D. Protoplast filtration and washing
1. Filter protoplast in a new Petri dish using a 60 μm nylon sieve.
2. Gently transfer filtered protoplast to a 15 mL Falcon.
3. Centrifuge the mixture at 100× g for 6–8 min.
4. Remove the supernatant.
5. Completely resuspend the protoplast pellet in 2–3 mL of wash solution.
6. Add wash solution until a final volume of 5–10 mL.
7. Centrifuge at 100× g for 3 min at 20 °C.
8. Remove the supernatant.
9. Repeat steps D5 to D8.
10. Resuspend protoplast pellet in 3–5 mL of wash solution.
E. Evaluation of protoplasts viability by fluorescein diacetate (FDA) staining
1. Add 0.5 mg/mL FDA solution to a previously obtained aliquot of protoplasts mixture.
2. Mix by pipetting.
3. Prepare a microscope slide and observe protoplasts (Figure 2G) under UV light (Figure 2H).
F. Protoplasts count, cultivation, and analysis of regenerative structures
1. Prepare a 1:10 protoplast dilution in wash solution and place an aliquot in a counting chamber.
2. Determine the number of protoplasts obtained in 1 mL (protoplast/mL) using an optical microscope.
3. Proceed to protoplast cultivation, using a final concentration of 1 × 105 ppt/mL.
4. Based on the yield obtained per milliliter, calculate the volume of suspension to obtain a quantity of 1 × 105 protoplasts and transfer it into 15 mL Falcons.
5. Centrifuge at 100× g for 3 min.
6. Remove the supernatant.
7. Resuspend each pellet in 1 mL of solid culture medium for protoplast cultivation and prepare four drops for each 60 mm Petri dish.
8. Once the drops have solidified, add 4 mL of liquid culture medium for protoplast cultivation.
9. Incubate plates at 27 °C.
10. Monitor cell divisions and development of regenerative structures every week using an inverted microscope (Figure 3A–F).
Figure 3. Stages of regeneration from protoplasts. A. First cell division (scale bars: 30 μm). B. Further cell divisions (scale bars: 40 μm). C. Somatic embryo at early globular stage of development (scale bars: 50 μm). D. Somatic embryo at the early globular stage of embryo development (scale bars: 70 μm). E. Somatic embryo at the heart stage of embryo development (scale bars: 70 μm). F. Somatic embryo at the torpedo stage of embryo development (scale bars: 100 μm).
Protoplasts PEG-mediated transfection
A. RNA guide design
RNA guide design was performed using CRISPOR online software (http://crispor.tefor.net) and CRISPR-RGENE (http://www.rgenome.net).
B. RNA guide synthesis
RNA guide synthesis was conducted using the GeneArt Precision gRNA Synthesis kit following the instructions provided by the manufacturer. The main steps are as follows:
1. Design forward and reverse oligonucleotides for PCR assembly.
2. PCR-assemble the gRNA DNA template using the PhusionTM High-Fidelity PCR master mix.
3. Generate the gRNA by in vitro transcription using the TranscriptAidTM enzyme mix.
4. Remove the DNA template by DNase I degradation.
5. Purify the in vitro–transcribed gRNA using the GeneJETTM purification columns.
6. Measure the purified gRNA concentration using Qubit RNA BR kit following the instructions provided by the manufacturer.
C. Cas9-sgRNA ribonucleoprotein complex assembly
To assemble the Cas9-sgRNA ribonucleoprotein complex, proceed as follows:
1. Add the Cas9 protein and sgRNA at a 1:1 weight ratio in a tube and mix gently.
2. Incubate for 10 min in darkness at room temperature.
D. PEG-mediated protoplasts transfection, cultivation, and analysis of regenerative structures
1. Add 2 × 105 protoplasts in a sterile 15 mL falcon tube.
2. Centrifuge the protoplasts at 100× g for 3 min.
3. Remove the supernatant.
4. Gently resuspend the protoplasts in 200 μL of MMG solution.
5. Add the Cas9-sgRNA ribonucleoprotein complex preassembled as described above in section “Cas9-sgRNA ribonucleoprotein complex assembly”.
6. Add 200 μL of PEG solution.
7. Mix gently until the solution is completely homogenized.
8. Incubate the solution for 20 min in the dark at room temperature.
9. Wash the protoplasts by adding 2 mL of sterile W5 solution and mix gently.
10. Centrifuge the protoplasts at 100× g for 3 min.
11. Remove the supernatant.
12. Repeat steps D9 to D11.
13. Remove the supernatant and resuspend the protoplasts in 1 mL of solid culture medium for protoplast cultivation. Proceed with the protoplasts cultivation as indicated in section above “Protoplasts count, cultivation, and analysis of regenerative structures” (steps F7 to F9).
14. Monitor cell divisions and the development of regenerative structures every week using an inverted microscope (Figure 3A–3F).
Somatic embryogenesis and plant regeneration
A. Identification of somatic cotyledonary embryos
1. After 2–3 months, verify the correct development of mature somatic cotyledonary embryos (Figure 4A) regenerated from protoplasts using a stereomicroscope.
Note: This step can be performed in non-sterile conditions.
Figure 4. Plants regeneration from protoplasts. A. Mature cotyledonary somatic embryo (scale bar: 0.5 mm). B. Germinated somatic embryo. C. Young plantlet in shooting medium. D. Whole plant regenerated in vitro.
B. Germination of somatic cotyledonary embryos
1. Using a stereomicroscope, transfer mature somatic cotyledonary embryos from DC medium to full germination medium.
2. Incubate for 3–4 weeks in darkness at 27 °C.
C. Shoots and plantlets development
1. Transfer well-developed germinated somatic embryos (Figure 4B) with forceps to C2D, C2D-4B, MG1, and MG1-10B media.
2. Incubate for 5–6 weeks under light with a 16-h photoperiod.
D. Plantlets regeneration and in vitro whole plant development
1. Check the correct development of regenerated plantlets (Figure 4C).
2. Transfer plantlets with forceps to MSN and RIM plant development media.
Note: Regenerated plantlets from C2D and C2D-4B were transferred to MSN medium, while regenerated plantlets from MG1 and MG1-10B were transferred to RIM medium.
3. Maintain fully regenerated plantlets (Figure 4D) under light with a 16-h photoperiod.
Validation of protocol
As mentioned in Najafi et al. 2022, to confirm genome editing events, genomic DNA was extracted from leaves of each regenerated plantlet derived from RNP-transfected protoplasts. A PCR amplification was performed targeting the specific genomic region of interest. Finally, Sanger sequencing verified mutations of the target site, confirming the success and precision of the editing process.
General notes and troubleshooting
General notes
1. Collect young unopened leaves from the apex of in vitro grapevine plantlets (5–6 weeks old) using sterile forceps. "Unopened" refers to leaves where the two sides of the leaf flap are still closed. Place droplets of sterile double-distilled H2O in a sterile Petri dish to maintain humidity.
2. Submerge inflorescences in water during processing and use two syringes to remove the calyptra, ensuring stamens remain attached to the pistil.
3. Embryogenic callus formation efficiency is dependent on the cultivar. Examine explants under a stereomicroscope for embryogenic material, which should appear white and granular. High-quality calli crumble apart when touched with a sterile scalpel.
4. After being placed in C1P medium, not all selected calli will proliferate effectively; use only high-quality material for subsequent steps.
5. Use cut tips to minimize damage to protoplasts and evaluate viability with fluorescein diacetate (FDA) staining. Monitor cell wall digestion and handle protoplasts carefully to maintain integrity.
6. The tissue of origin for protoplast isolation, details in the purification steps, and the composition of the regeneration media may vary across plant species, whereas the type and concentration of enzymes in the digestion solution are quite standard regardless of the starting material.
Acknowledgments
E.B., E.D. and S.Ze. designed the research; E.B., S.Za., S.L., L.C. and C.C. performed the research and analyzed the data; A.A., M.F. and S.Ze. contributed to data analysis; E.B., E.D., S.Za., S.L., L.C. and C.C. wrote the paper. This work was supported by EdiVite s.r.l., Grant BAYER (University of Verona awarded to Sara Zenoni) and PRIN 2022 (University of Verona awarded to Sara Zenoni). This protocol is based on our recent publication [13].
Competing interests
The patent application N. 102021000023468 is based on this protocol.
References
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Article Information
Publication history
Received: Jul 19, 2024
Accepted: Dec 4, 2024
Available online: Jan 9, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Category
Plant Science > Plant cell biology > Cell isolation
Plant Science > Plant molecular biology > Genetic analysis
Biological Sciences > Biological techniques > CRISPR/Cas9
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Precise Generation of Human Induced Pluripotent Stem Cell–derived Cell Lines Harboring Disease-relevant Single Nucleotide Variants Using a Prime Editing System
SK Seiya Kanno
KS Kota Sato
TN Toru Nakazawa
In Press, Available online: Jan 05, 2025
DOI: 10.21769/BioProtoc.5191 Views: 34
Reviewed by: Philipp WörsdörferWei Dai Anonymous reviewer(s)
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Abstract
Human induced pluripotent stem (iPS) cell lines harboring mutations in disease-related genes serve as invaluable in vitro models for unraveling disease mechanisms and accelerating drug discovery efforts. Introducing mutations into iPS cells using traditional gene editing approaches based on the CRISPR-Cas9 endonuclease often encounters challenges such as unintended insertions/deletions (indels) and off-target effects. To address these limitations, we present a streamlined protocol for introducing highly accurate gene mutations into human iPS cells using prime editing, a “search-and-replace” genome-editing technology that combines unwanted indel-minimized CRISPR-Cas9 nickase with reverse transcriptase. This protocol encompasses the design of prime editing guide RNAs (pegRNAs) required for binding and replacement at target loci, construction of prime editor and pegRNA expression vectors, gene transfer into iPS cells, and cell line selection. This protocol allows for the efficient establishment of disease-associated gene variants within 6–8 weeks while preserving critical genomic context.
Key features
• Dramatic improvement in efficiency of In-Fusion cloning using inserts assembled from the three pegRNA components (spacer, spCas9 scaffold, and 3' extension) via overlap extension PCR.
• Cost-effective and time-saving selection of pegRNAs for prime editing via bulk Sanger sequencing.
• Straightforward gene transfection using polymer-based reagents, which requires no specialized equipment or techniques and offers high reproducibility and broad applicability across different cell lines.
• Precise genome editing based on pegRNA/prime editing minimizes off-target effects, enabling a wide range of applications in the study of disease-associated genetic variants.
Keywords: Human induced pluripotent stem (iPS) cell Prime editing CRISPR/Cas9 Precision genome editing pegRNA design Disease-associated single nucleotide variant (SNV) Disease modeling Isogenic cell line
Graphical overview
Key steps of generation of human induced pluripotent stem (iPS) cell lines harboring disease-relevant single nucleotide variants (SNVs) using a prime editing system.
Background
Advances in genomic disease analysis have identified numerous single nucleotide variants (SNVs) associated with diverse diseases. Patient-derived human induced pluripotent stem (iPS) cells carrying these SNVs offer valuable in vitro models for elucidating disease mechanisms and accelerating drug discovery. However, phenotypic variability arising from diverse genetic backgrounds among iPS cell lines can confound the accurate assessment of disease gene effects [1].
Recent advancements in genome editing technologies have enabled the establishment of highly improved protocols for generating isogenic iPS cell lines through homology-directed repair (HDR)-mediated SNVs editing using CRISPR-Cas9 and single-stranded oligodeoxynucleotides (ssODNs), facilitating functional analyses of SNVs [2–5]. Nonetheless, these methods often introduce unintended insertions or deletions (indels) and off-target effects due to the induction of double-strand breaks in the DNA by CRISPR-Cas9, limiting their precision and efficiency [6].
To address these limitations, we present a protocol for generating iPS cell lines with precisely engineered SNVs in genes of interest using prime editing. Prime editing is a “search-and-replace” genome editing tool that combines a modified CRISPR-Cas9 nickase with reverse transcriptase to minimize unwanted indels [6]. This approach enables the functional analysis of gene variants while preserving the genomic context, providing a more controlled and informative system for studying disease mechanisms. Specifically, this protocol presents a workflow for generating a cell line harboring the OPTN p.(Asn51Thr) missense mutation, which was previously identified in our laboratory as an SNV associated with normal-tension glaucoma (NTG) [7].
This protocol has several key steps. First, the necessary components for the prime editing system are prepared, including the prime editor enzyme and prime editing guide RNA (pegRNA) expression vectors designed to introduce specific mutations. Additionally, a selectable marker gene, such as the puromycin resistance gene, is included to identify and select cells that have successfully taken up the editing components. Next, each pegRNA is introduced into HEK293T cells along with the prime editor enzyme. To assess the efficiency of the designed pegRNAs, the sequence of the target loci is analyzed in a bulk population of transfected cells. Finally, the pegRNA and prime editor enzyme are introduced into a human iPS cell line, 201B7, and individual iPS cell clones are isolated from the transfected population. By focusing on pegRNAs with high editing efficiency, the chances of obtaining iPS cell lines with the desired gene mutations are increased.
This protocol offers a robust and efficient method for generating iPS cell lines with targeted gene mutations, not only for studying disease-related genes but also for correcting pathogenic variants in patient-derived cells. While prime editing offers significant advantages over traditional methods, it is not without its limitations. For example, although several efficient pegRNA design tools are available, more information is needed in the future regarding editing efficiency in human iPS cells, as it can vary depending on the target locus and cell type. Furthermore, experimental conditions, such as reagent concentrations, transfection methods, and selection markers, must be carefully optimized for each iPS cell line to achieve optimal results.
Materials and reagents
Biological materials
1. Competent quick DH5α (Toyobo, catalog number: DNA-913F); store in a -80 °C ultra-low temperature freezer
2. Human iPS cell line 201B7 (RIKEN BRC Cell Bank, catalog number: HPS0063); to prevent a decrease in cell viability, iPS cells should be immediately transferred to liquid nitrogen for storage after receiving them from the supplier or after creating a cell bank
3. pLV[Exp]-EF1A>hCas9(ns):T2A:Puro (VectorBuilder, catalog number: VB210412-1054sbc)
4. pCMV-PEmax-P2A-hMLH1dn (Addgene, catalog number: 174828)
5. pU6-pegRNA-GG-acceptor (Addgene, catalog number: 132777)
6. pMK232 (CMV-OsTIR1-PURO) (Addgene, catalog number: 72834)
Store plasmids in a freezer at -30 °C.
Reagents
1. StemFit medium (Ajinomoto Healthy Supply, catalog number: AK03N)
2. DMEM (4.5 g/L glucose) with L-Gln and sodium pyruvate (Nacalai Tesque, catalog number: 08458-45)
3. Fetal bovine serum (FBS) (Thermo Fisher Scientific, catalog number: 12483020)
4. DPBS, no calcium, no magnesium (Thermo Fisher Scientific, catalog number: 14190144)
5. Trypsin-EDTA (0.25%), phenol red (Thermo Fisher Scientific, catalog number: 25200056); dilute with DPBS as needed.
6. TrypLE Select enzyme (1×), no phenol red (Thermo Fisher Scientific, catalog number: 12563011)
7. Penicillin-streptomycin (Thermo Fisher Scientific, catalog number: 15140122)
8. CultureSure Y-27632 (Fujifilm Wako Pure Chemical, catalog number: 036-24023)
9. iMatrix-511 (Matrixome, catalog number: 892012)
10. STEM-CELLBANKER GMP grade (Zenogen Pharma, catalog number: 11924)
11. PolyJet DNA in vitro transfection reagent (SignaGen Laboratories, catalog number: SL100688)
12. Puromycin (InvivoGen, catalog number: ant-pr)
13. QIAamp DNA Mini kit (Qiagen, catalog number: 51306)
14. NucleoSpin gel and PCR clean-up (Macherey-Nagel, catalog number: 740609.250)
15. SOC medium (Thermo Fisher Scientific, catalog number: 15544034)
16. Plasmid Miniprep Plus Purification kit (BioElegen Technology, catalog number: DP01-PLUS-300)
17. KOD One PCR Master Mix (Toyobo, catalog number: KMM-101)
18. Dpn I (New England Biolabs, catalog number: R0176L)
19. In-fusion snap assembly master mix (Takara Bio, catalog number: 638947)
20. Ampicillin sodium (Fujifilm Wako Pure Chemical, catalog number: 016-23301)
21. BD Difco LB (Luria-Bertani) broth Miller (Becton, Dickinson and Company, catalog number: 244620)
22. BD BACTO agar (Becton, Dickinson and Company, catalog number: 214010)
23. BD Difco 2×YT (yeast extract tryptone medium) (Becton, Dickinson and Company, catalog number: 244020)
24. Agarose S (Nippon Gene, catalog number: 318-01195)
25. Gel loading dye, purple (6×), no SDS (New England Biolabs, catalog number: B7025S)
26. 1 kb DNA ladder (Nippon Genetics, catalog number: NE-MWD1P)
27. 100 bp DNA ladder (Takara Bio, catalog number: 3422D)
28. Primers (OPC cartridge purified from Eurofins Genomics)
Homologous sequences for overlap PCR or in-fusion cloning are displayed in lowercase:
OPTN_genotyping_forward: 5'- AATCGCCAATGGGTTTGTGGGAC -3'
OPTN_genotyping_reverse: 5'- TGCTAAATCCTGTGCTTCCCCACC -3'
EF1A_ forward: 5'- ccagatatacgcgttGGCTCCGGTGCCCGTCAGTG -3'
EF1A_reverse: 5'- acggttcactaaaccTCACGACACCTGAAATG -3'
Inverse_PEmax_forward: 5'- ggtttagtgaaccgtCAGATCCGC -3'
Inverse_PEmax_reverse: 5'- aacgcgtatatctggCCCGTACATC -3'
EF1A_sequence_forward1: 5'- GGTGGAGACTGAAGTTAGGCCAGC -3'
EF1A_sequence_forward2: 5'- TAAGTGCAGTAGTCGCCGTGAACG -3'
EF1A_sequence_reverse: 5'- CACGCAAGGGCCATAACCCG -3'
tevopreQ1+Terminator_forward: 5'- CGCGGTTCTATCTAGTTACGCGTTAAACCAACTAGAA -3'
tevopreQ1+Terminator_reverse: 5'- AAAAAATTCTAGTTGGTTTAACGCGTAACTAGATAGAACCGCG -3'
Spacer_OPTN_forward: 5'- gaaaggacgaaacaccGCTGCTCACCTTTCAGCTGG -3'
Spacer_OPTN_reverse: 5'- ttctagctctaaaacCCAGCTGAAAGGTGAGCAGC -3'
spCas9_scaffold_forward: 5'- GTTTTAGAGCTAGAAATAGCAAGTTAAAATAAGGCTAGTCCGTTATCAACTTGA -3'
spCas9_scaffold_reverse: 5'- GCACCGACTCGGTGCCACTTTTTCAAGTTGATAACGGACTAGCCTTATTTTAACTTGC -3'
3’extension+Linker_OPTN_forward: 5'- gcaccgagtcggtgcTGACCGAGACCCACCAGCTGAAAGGACCC -3'
3’extension+Linker_OPTN_reverse: 5'- ctagatagaaccgcgATTAGGGTCCTTTCAGCTGGTGGGTCTCGGTC -3'
Inverse_U6-pegRNA_Forward: 5'- cgcggttctatctagTTACGCG -3'
Inverse_U6-pegRNA_reverse: 5'- ggtgtttcgtcctttcCACAAGA -3'
pegRNA_sequence_forward: 5'- GAGGGCCTATTTCCCATGATTCC -3'
Laboratory supplies
1. 15 mL centrifuge tubes (TPP, catalog number: 91015)
2. 1.5 mL microcentrifuge tubes (Ina-optika, catalog number: 113004)
3. 0.5 mL microcentrifuge tube (Ina-optika, catalog number: SC-005)
4. 20 μL racked tips, sterile (Eppendorf, catalog number: 0030075226)
5. 200 μL racked tips, sterile (Eppendorf, catalog number: 0030075234)
6. 1000 μL racked tips, sterile (BM Equipment, catalog number: WE1000-RL)
7. 12-well cell culture plate (Corning, catalog number: 353043)
8. 24-well cell culture plate (Corning, catalog number: 353047)
9. 48-well cell culture plate (Corning, catalog number: 353078)
10. 96-well cell culture plate (Corning, catalog number: 353072)
11. 60 × 15 mm cell culture dish (Corning, catalog number: 353004)
12. STAR SDish9015 ver. 2 Petri dish (Rikaken, catalog number: RSU-SD9015-2)
13. 0.2 mL 8-tube PCR strips (Bio-Rad Laboratories, catalog number: TLS0801)
14. 0.2 mL domed PCR tube 8-cap strips (Bio-Rad Laboratories, catalog number: TCS0801)
15. 2 mL cryo vial, inner screw (Simport Scientific, catalog number: T311-2)
16. BICELL freezing treatment container (Nihon Freezer, catalog number: BICELL)
Equipment
1. Thermal cycler (Bio-Rad, model: C1000 Touch)
2. Refrigerated centrifuge (Tomy Seiko, model: LX-120)
3. High-speed refrigerated microcentrifuge (Tomy Seiko, model: MX300)
4. Cell culture incubator (PHC, catalog number: MCO-170AICUVD-PJ)
5. Shaker incubator (Taitec, model: BR-40LF)
6. Benchtop incubator for bacterial culture (AS ONE, model: PIC-100)
7. Microvolume spectrophotometer (Thermo Fisher Scientific, model: NanoDrop 2000c)
8. Gel electrophoresis equipment (Takara Bio, model: Mupid-exU)
9. Gel imaging system (Bio-Rad, model: GelDoc Go imaging system)
10. Autoclave (Tomy Seiko, model: LBS-245)
Software and datasets
1. SnapGene Viewer (SnapGene, https://www.snapgene.com/snapgene-viewer)
Procedure
A. Identification of the genotype at the targeted SNV locus in the cell line
As the first step, it is necessary to investigate the impact of the target gene mutation in advance, as the effort required to establish cell lines differs depending on whether heterozygous or homozygous cell lines are ultimately needed after editing. Databases such as ClinVar (https://www.ncbi.nlm.nih.gov/clinvar/) and gnomAD (https://gnomad.broadinstitute.org/) can be used to search for information about the target mutation and reports from other researchers. For example, missense mutations like OPTN p.(Glu50Lys), which cause early-onset familial NTG, often exhibit pathogenicity even in a heterozygous state in autosomal-dominant genetic diseases. Additionally, mutations with haploinsufficiency or a dominant negative effect are also likely to be pathogenic in a heterozygous state, so depending on the research purpose, a heterozygous cell line may be sufficient.
Next, it is necessary to identify the genotype at the target disease-related locus in the cell line to be used. If the genotype of the cell line to be used is heterozygous, pegRNAs that convert the wild type and mutant types to each other (wild type to mutant type or vice versa) are designed, and prime editing is used with each pegRNA in separate experiments to obtain homozygous cell lines of both the wild type and mutant types. On the other hand, if starting from a homozygous genotype, it is necessary to increase the number of single clones to be obtained in section E, as the frequency of obtaining homozygous cell lines after editing is lower than that of heterozygous cell lines. Alternatively, it is possible to obtain homozygous cell lines by performing editing again after obtaining a heterozygous cell line in a single editing process.
1. Retrieve the genomic sequence surrounding the variant of interest.
a. Search for the gene of interest (e.g., OPTN) in the gnomAD database (https://gnomad.broadinstitute.org/).
b. Locate the variant ID of the target variant. If unavailable, use a nearby variant’s ID (e.g., OPTN p.Glu50Lys for OPTN p.Asn51Thr; not listed).
c. Follow the UCSC link under “External Resources” to visualize the genomic region surrounding the variant.
d. Download the genomic sequence in FASTA format from the “DNA sequence” link under the View menu in UCSC.
e. Store the downloaded sequence in a sequence analysis tool such as SnapGene Viewer.
2. Design genotyping primers.
a. Paste the target locus nucleotide sequence into the PCR template box of Primer-BLAST (https://www.ncbi.nlm.nih.gov/tools/primer-blast/).
b. Set primer design parameters: amplicon length of 500–700 bp, Tm of 62 °C ± 3 °C, GC content of 50%–80%, and a GC clamp of 1. Click the Get Primers button to search for potential primer pairs.
c. Order primers from a qualified supplier (e.g., Eurofins Genomics).
Note 1: When ordering from Eurofins Genomics, order a 10-nmol-scale oligonucleotide OPC cartridge purified and dissolved in a TE buffer. Upon arrival, store at -20 °C and thaw immediately before use.
Note 2: It is recommended to order at least three pairs of primers and conduct PCR experiments to identify the optimal primer pair that specifically amplifies the target gene without any off-target amplification.
3. Extract and quantify genomic DNA.
a. Extract genomic DNA from the cell line following the manufacturer’s instructions for the genomic DNA extraction kit (e.g., QIAamp DNA Mini Kit).
b. Quantify the extracted DNA concentration using a NanoDrop spectrophotometer at A260.
c. Store the DNA at -20 °C for up to 6 months.
4. Perform PCR with primers spanning the target to amplify and enrich the genomic region of interest.
a. Prepare a 20 µL PCR reaction mixture for each sample (Table 1).
To amplify the region of the OPTN p.(Asn51Thr) missense mutation, perform PCR using OPTN_genotyping_forward and OPTN_genotyping_reverse primers.
Table 1. PCR reaction mixture components
Reagent Final concentration Volume
KOD One PCR master mix (dye-free 2×PCR master mix) 1× 10 μL
Forward primer (2 μM) 0.3 μM 3 μL
Reverse primer (2 μM) 0.3 μM 3 μL
Genomic DNA (20 ng/µL) 4 ng/µL 4 μL
Total n/a 20 μL
b. Perform two-step PCR using a thermal cycler under the temperature conditions provided in Table 2.
Table 2. Thermocycling conditions for PCR
Step Temp. (°C) Duration No. of cycles
Initial denaturation 98 30 s 1
Denaturation 98 10 s 35
Extension 68 5 s/kb
Hold 4 ∞ -
c. Apply 1–2 μL of the PCR product to a 1.5% agarose gel containing a fluorescent nucleic acid stain (e.g., GelRed) and run electrophoresis at 100 V for approximately 15 min.
d. Visualize DNA bands on a transilluminator and verify that the PCR amplicon has migrated to the expected band size.
Note: Consider using touchdown PCR or nested PCR when you need to increase DNA yield or specificity, such as when amplifying low-abundance targets or dealing with non-specific amplification.
e. Purify the target amplicon DNA from PCR products using a commercial DNA purification kit (e.g., NucleoSpin Gel and PCR clean-up).
Note: For single bands, direct purification of DNA from the post-PCR mixture is possible. If extra bands are present, indicating non-specific amplification, excise the desired-sized band from the gel after electrophoresis and extract DNA from the gel.
f. Quantify the extracted DNA concentration using a NanoDrop spectrophotometer at A260. Store the DNA at -20 °C for up to 6 months.
5. Identify the genotype at the target disease-related locus in the cell line.
a. Send the PCR product to a sequencing service provider (e.g., Eurofins Genomics) for Sanger sequencing to determine the nucleotide sequence. PCR primers within 100–600 bp of the target locus can often serve as sequencing primers. Use OPTN_genotyping_forward as a sequencing primer for the region of the OPTN p.(Asn51Thr) missense mutant.
b. Input the nucleotide sequence obtained from Sanger sequencing into Standard Nucleotide BLAST (https://blast.ncbi.nlm.nih.gov/Blast.cgi?PROGRAM=blastn&BLAST_SPEC=GeoBlast&PAGE_TYPE=BlastSearch) to confirm the genotype of the cell line.
c. To account for potential heterozygosity, examine not only the FASTA sequence obtained from Sanger sequencing but also the waveform data. Check whether there are overlapping peaks of two different bases at the same position in the waveform data (Figure 1).
Figure 1. Identification of genotype by analyzing Sanger sequencing waveforms. The red box indicates the position of the OPTN p.(Asn51Thr) missense mutation. In cell lines with the wild-type genotype, this position exhibits a homozygous C/C genotype. In cell lines harboring the OPTN p.(Asn51Thr) missense mutation, this position displays either a homozygous A/A or a heterozygous C/A genotype. Heterozygosity for C/A is characterized by overlapping peaks of the two bases, often accompanied by lower-quality values.
B. Design of pegRNAs
PegRNAs are a pivotal component of the prime editing system, directing the targeted insertion of new DNA sequences into a specific genomic locus. A pegRNA consists of three primary domains: a spacer, a reverse transcriptase template (RTT), and a primer binding site (PBS). The spacer, in a complex with the Cas9 protein, recognizes and binds to the target DNA sequence, specifying the precise location for editing. The RTT serves as a template for reverse transcriptase, encoding the new DNA sequence to be inserted. The PBS determines the starting point for reverse transcription, ensuring accurate editing.
Engineered pegRNA (epegRNA), incorporating a trimmed, modified preQ1 riboswitch aptamer (tevopreQ1) at its 3' end, has become prevalent. The stable stem-loop structure formed by tevopreQ1 shields the epegRNA from degradation by cellular RNases, enhancing its stability and, consequently, prime editing efficiency [8]. A linker sequence is inserted between tevopreQ1 and the PBS to prevent unintended base pairing interactions.
In this step, the spacer, RTT, and PBS will be designed for the pegRNA. The design of pegRNA significantly influences both editing efficiency and specificity. Factors such as spacer length, RTT sequence, and PBS position must be optimized. To facilitate efficient design, various browser-based tools, including DeepPrime, pegFinder, and PE-Designer, are employed to generate optimal spacer, RTT, and PBS sequences. DeepPrime, for instance, leverages machine learning to design highly efficient pegRNAs with minimal off-target effects. As the next step of pegRNA design, an appropriate linker sequence is determined to avoid base pairing with other domains of the pegRNA and to ensure optimal reverse transcription. An RNA secondary structure prediction tool, pegLIT, is used for linker design.
1. Use pegRNA design tools such as DeepPrime (https://deepcrispr.info/DeepPrime/), pegFinder (http://pegfinder.sidichenlab.org/), and PE-Designer (http://www.rgenome.net/pe-designer/) to input the target genomic sequence and generate candidate spacer, RTT, and PBS sequences. These tools use machine learning or algorithmic approaches to design optimal pegRNAs.
Note 1: To optimize prime editing efficiency, it is recommended to pair an 11 nt PBS with an RTT of 12 nt or less, and a 12 nt PBS with an RTT of 13 nt or more [9].
Note 2: Regularly check for updates to pegRNA design tools, as features and functionalities may evolve.
2. Use RNA secondary structure prediction tools like pegLIT (https://peglit.liugroup.us/) to design optimal linker sequences based on the spacer, RTT, and PBS sequences.
3. Order custom-synthesized single-stranded oligonucleotides corresponding to the top and bottom strands of the spacer, 3' extension, linker, and tevopreQ1 sequences from a qualified supplier (e.g., Eurofins Genomics).
Critical: To initiate transcription from the RNA polymerase III–dependent U6 promoter, add a guanine or adenine nucleotide to the 5' end of the guide RNA sequence [10].
Critical: Order the oligonucleotides so that the 15 bases at the end of each DNA fragment [spacer, scaffold, 3' extension sequence (PBS + RTT + Linker), and backbone vector] are homologous to each other to allow for overlap PCR and in-fusion cloning (Figure 2). While the synthesis may be more costly and complex, DNA fragments spanning the spacer-linker region of each designed pegRNA can alternatively be used as inserts for in-fusion cloning.
Note 1: It is recommended to order at least three distinct pegRNA sequences and comparing their actual editing efficiencies to identify the most effective candidate.
Note 2: When ordering from Eurofins Genomics, order a 10-nmol-scale oligonucleotide OPC cartridge purified and dissolved in TE buffer. Upon arrival, store at -20 °C and thaw immediately before use.
Figure 2. Designed guide RNA for introducing the p.(Asn51Thr) point mutation into the OPTN gene using prime editing. PCR primers targeting the spacer and 3' extension+linker are flanked by 15 bp or longer homology regions to enable efficient assembly by overlap PCR or in-fusion cloning.
C. Preparation of constructs for prime editing
Prime editing (PE) systems have undergone significant advancements in recent years. Initial PE1 systems, which fused SpCas9 with Moloney murine leukemia virus (M-MLV) reverse transcriptase (RT), exhibited limited genome editing efficiency in human cells [11]. PE2 used engineered RT, while PE3 further improved efficiency by introducing a nick into the complementary strand using an additional sgRNA [6]. PEmax is a prime editor protein with optimized codons for both RT and Cas9. PE4 and PE5 transiently inhibited DNA mismatch repair (MMR) by using a dominant-negative mutant of MLH1, a key factor in the MMR pathway, thereby improving editing efficiency. The combination of these systems in PE4Max and PE5Max achieved even higher editing efficiencies [12]. Additionally, the PE7 system improved the stability and integrity of the pegRNA by fusing the N-terminal domain of the RNA-binding protein La to the prime editor protein, thereby enhancing editing efficiency. [13].
This step involves the construction of two essential plasmid vectors for the prime editing system (Figure 3). Each plasmid vector is composed of multiple components that need to be assembled to execute this protocol. Specifically, these include the prime editor gene, pegRNA sequence, and puromycin resistance gene for the selection of transfected cells. For this protocol, we used pCMV-PEmax-P2A-hMLH1dn (Addgene, plasmid number: 174828) as the PE4max expression vector. For next-generation PE7 [11], pCMV-PE7 (Addgene, plasmid number: 214812) should be purchased. In pluripotent stem cells, selecting a constitutive promoter that ensures robust gene expression is crucial. Previous studies have shown that the transcriptional activity of the CMV enhancer is significantly lower than that of the EF1A or CAG promoter in both human [14] and mouse [15] pluripotent stem cells. Therefore, we recommend replacing the CMV enhancer in pCMV-PEmax-P2A-hMLH1dn with the EF1A promoter to drive prime editor expression more effectively. Plasmids constructed using a similar strategy, including an EF1A promoter–driven prime editor and a pegRNA backbone vector with a puromycin resistance gene, are available from Addgene (plasmid numbers 184444 and 214085, respectively).
Figure 3. Components of the plasmid vector required for the prime editing system. A. A plasmid vector co-expressing prime editor and MLH1dn under the EF1A promoter. The prime editor protein is a fusion protein comprising SpCas9 nickase (nCas9) and Moloney murine leukemia virus (M-MLV) reverse transcriptase (RT). B. A plasmid vector expressing pegRNA under the U6 promoter and a puromycin resistance gene under the PGK promoter. The pegRNA consists of a spacer sequence specific to the target gene, a scaffold sequence required for complex formation with spCas9, a 3' extension sequence (labeled “3'Ext”) for editing, a linker sequence (labeled “L”), a tevopreQ1 sequence for enhancing RNA stability, and a terminator sequence (labeled “T”).
1. Amplify the insert fragment and backbone vector with PCR to obtain sufficient amounts for subsequent in-fusion cloning.
a. Prepare a 20 μL PCR reaction mixture (Table 3).
To amplify the EF1A promoter region, perform PCR using pLV[Exp]-EF1A>hCas9(ns):T2A:Puro (containing the EF1A promoter) as a template and EF1A_forward and EF1A_reverse primers.
To amplify the backbone, perform PCR using pCMV-PEmax-P2A-hMLH1dn as a template and Inverse_PEmax_forward and Inverse_PEmax_reverse primers.
Table 3. PCR reaction mixture components
Reagent Final concentration Volume
KOD One PCR master mix (dye-free 2×PCR master mix) 1× 10 μL
Forward primer (2 μM) 0.3 μM 3 μL
Reverse primer (2 μM) 0.3 μM 3 μL
Plasmid DNA (5 ng/µL) 1 ng/µL 4 μL
Total n/a μL
b. Perform two-step PCR using a thermal cycler under the temperature conditions provided in Table 4.
Table 4. Thermocycling conditions for PCR
Step Temp. (°C) Duration No. of cycles
Initial denaturation 98 30 s 1
Denaturation 98 10 s 35
Extension 68 5 s/kb
Hold 4 ∞ -
c. Apply 1–2 μL of the PCR product to a 1.5% agarose gel containing a fluorescent nucleic acid stain (e.g., GelRed) and run electrophoresis at 100 V for approximately 15 min.
d. Visualize DNA bands on a transilluminator and verify that the PCR amplicon has migrated to the expected band size.
Note: Consider using touchdown PCR or nested PCR when you need to increase DNA yield or specificity, such as when amplifying low-abundance targets or dealing with non-specific amplification.
e. To remove the parental vector template, add 5–10 units of DpnI to 20 μL of the PCR post-reaction mixture and incubate at 37 °C for 10 min. Then, deactivate the enzyme by incubating at 80 °C for 10 min.
Note: Plasmid DNA amplified in common Escherichia coli strains is methylated, while PCR products are typically unmethylated. DpnI specifically cleaves plasmids with methylated recognition sites. Therefore, linearized plasmids amplified by PCR using plasmid DNA as a template remain intact, while the template plasmid is degraded, preventing its carryover into transformation.
f. Purify the target amplicon DNA from PCR products using a commercial DNA purification kit (e.g., NucleoSpin gel and PCR clean-up).
Note 1: For single bands, direct purification of DNA from the post-PCR mixture is possible. If extra bands are present, indicating non-specific amplification, excise the desired-sized band from the gel after electrophoresis and extract DNA from the gel.
Note 2: Extensive washing twice with the wash buffer of the DNA purification kit has been shown to enhance the efficiency of subsequent in-fusion cloning.
g. Quantify the extracted DNA concentration using a NanoDrop spectrophotometer at A260. Store the DNA at -20 °C for up to 6 months.
2. Mix the PCR products from step C1, the insert fragment, and the linearized vector, and perform cloning using the in-fusion snap assembly master mix.
a. Prepare a 5 μL in-fusion cloning reaction mixture. Mix the insert fragment and the linearized plasmid vector so that the approximate molar ratio is 2:1 (Table 5).
Note: Carry out an in-fusion cloning reaction without the inclusion of an insert fragment to serve as a negative control for subsequent transformation experiments.
Table 5. In-fusion cloning reaction mixture components
Reagent Final concentration Volume
In-fusion snap assembly master mix (5×) 1× 1 μL
Insert fragment 1–20 ng/μL 1 μL
linearized plasmid vector 5–20 ng/μL 1 μL
Nuclease-free water n/a To 5 μL
Total n/a 5 μL
b. Carry out the in-fusion cloning reaction in a thermal cycler using the temperature conditions provided in Table 6.
Table 6. Thermocycling conditions for PCR
Step Temp. (°C) Duration
In-fusion reaction 1 37 15 min
In-fusion reaction 2 50 15 min
Hold 4 ∞
c. Store the cloning reactions at -20 °C until ready to proceed with the transformation.
3. Transform E. coli with plasmids constructed using in-fusion cloning.
a. Combine 20 μL of competent cells with 0.5 μL of the in-fusion cloning reaction mixture and incubate on ice for 2 min.
Critical: To ensure a successful transformation, it is recommended that the amount of DNA is less than 2 ng for 20 μL of competent cells, and that the volume of TE buffer provided does not exceed 30% of the volume of the competent cells.
Note: Use the in-fusion cloning mixture lacking the insert fragment from step C2a as a negative control for transformation.
b. Heat shock the cells at 42 °C for 30 s and immediately transfer the cells to ice for 2 min.
c. Add 50 μL of SOC medium to the cells and shake at 37 °C for 30 min.
d. Spread the transformation mixture onto LB agar plates containing 40 µg/µL of ampicillin.
e. Incubate at 37 °C overnight (12–16 h).
Note: If a high number of background colonies are observed on a negative control plate, this may indicate incomplete DpnI digestion and, consequently, reduced in-fusion cloning efficiency. To address this, increase the amount of DpnI added or extend the DpnI enzymatic reaction time.
4. Perform colony PCR to select an E. coli colony that has been transformed with a plasmid vector into which the insert fragment has been inserted.
a. Prepare a 10 μL colony PCR reaction mixture (Table 7).
To amplify the EF1A promoter region, perform PCR using EF1A_forward and EF1A_reverse primers.
Table 7. Colony PCR reaction mixture components
Reagent Final concentration Volume
KOD One PCR master mix (dye-free 2× PCR master mix) 1× 10 μL
Forward primer (2 μM) 0.3 μM 3 μL
Reverse primer (2 μM) 0.3 μM 3 μL
Nuclease-free water n/a To 5 μL
b. Using a sterile pipette tip, transfer a small amount of each of 6–10 individual well-formed colonies to separate PCR tubes containing the prepared PCR mixture.
Note: Use a permanent marker to number each colony on the plate to correlate the colony PCR results with the corresponding colonies.
c. Perform two-step PCR using a thermal cycler under the temperature conditions in Table 8.
Table 8. Thermocycling conditions for PCR
Step Temp. (°C) Duration No. of cycles
Initial denaturation 98 30 s 1
Denaturation 98 10 s 30
Extension 68 5 s/kb
Hold 4 ∞ -
d. Apply 1–2 μL of the PCR product to a 1.5% agarose gel containing a fluorescent nucleic acid stain (e.g., GelRed) and run electrophoresis at 100 V for approximately 15 min.
e. Visualize DNA bands on a transilluminator to identify colonies transformed with the successfully in-fusion-cloned plasmid vector.
5. Culture the transformed E. coli in liquid medium.
a. Using a pipette tip, touch the surface of 2–3 colonies confirmed to be positive for the insert fragment by colony PCR and inoculate them into 10 mL of BD DIFCO 2×YT (yeast extract tryptone broth) containing 40 µg/mL ampicillin in a 50 mL centrifuge tube.
Note: To prevent cross-contamination, use a fresh pipette tip for each colony.
b. Incubate at 37 °C for 16–20 h in a shaking incubator at 120–150 rpm.
Note: To promote aerobic respiration in E. coli, slightly loosen the cap to improve aeration and tilt the tube slightly.
c. Purify plasmids propagated by E. coli using commercial Miniprep kits (e.g., Plasmid Miniprep Plus Purification kit, BioElegen Technology).
Critical: Use an endotoxin-free plasmid purification kit to avoid unexpected cellular responses, such as cytotoxicity, during transfection.
d. Quantify the plasmid DNA concentration using a NanoDrop spectrophotometer at A260. Store the circular plasmids at -20 °C for up to 6 months.
Critical: Ensure the concentration of the purified plasmid DNA is at least 1,000 ng/µL to avoid negative impacts on cell viability and transfection efficiency, which can occur when excessive elution buffer is used.
6. Confirm the absence of DNA base substitutions around the in-fusion recombination site in the constructed plasmid vector with Sanger sequencing. Use EF1A_sequence_forward1, EF1A_sequence_forward2, and EF1A_sequence_reverse as sequencing primers.
7. Construct the pegRNA expression vector using PCR amplification and in-fusion cloning, following the same procedure as in steps C1–5. The backbone of the pegRNA expression vector, excluding sequence spanning from spacer to linker, is common regardless of the objective genomic locus or mutation. Clone this common sequence (e.g., tevopreQ1 and puromycin resistance gene) to create a backbone construct. Subsequently, clone the variable pegRNA sequences into the backbone construct using in-fusion cloning.
Note: While it is possible to directly insert three fragments (spacer, scaffold, and 3' extension+linker) into a vector using in-fusion cloning, we found that this approach often results in low cloning efficiency, typically less than 10%. To enhance efficiency, we recommend pre-assembling these fragments using overlap PCR (Figure 4). This method has consistently yielded cloning efficiencies of over 95% in our experiments. Although it requires additional synthesis costs, another option is to use a continuous DNA fragment spanning the entire region from the spacer to the linker, tailored to each designed pegRNA sequence.
a. Amplify the insert fragment and backbone vector with PCR, following the same procedure as in steps C1a–d.
To amplify the spacer, spCas9 scaffold, 3’ extension+Linker sequence, and backbone, perform PCR using the corresponding primer pairs: Spacer_OPTN_forward/reverse, spCas9_scaffold_forward/reverse, 3’extension+Linker_OPTN_forward/reverse, and Inverse_U6-pegRNA_forward/reverse, respectively.
To remove the parental vector template, add 5–10 units of Dpn I to 20 μL of the PCR post-reaction mixture and incubate at 37 °C for 10 min, then deactivate the enzyme by incubating at 80 °C for 10 min.
b. Perform overlap PCR using the PCR-amplified spacer, spCas9 scaffold, and 3’extension+linker fragments as templates to obtain the desired pegRNA sequence as the PCR product. Use primers that amplify from both ends of the assembled pegRNA. Prepare a 20 μL PCR reaction mixture for each sample (Table 9).
To amplify the pegRNA sequence, perform PCR using Spacer_OPTN_forward and 3’extension+Linker_OPTN_reverse primers.
Table 9. Overlap PCR reaction mixture components
Reagent Final concentration Volume
KOD One PCR master mix (dye-free 2×PCR master mix) 1× 10 μL
Forward primer (2 μM) 0.3 μM 3 μL
Reverse primer (2 μM) 0.3 μM 3 μL
Post-PCR mixture (spacer) n/a 1 μL
Post-PCR mixture (spCas9 scaffold) n/a 2 μL
Post-PCR mixture (3’extension+Linker) n/a 1 μL
Total n/a 20 μL
c. Perform PCR following the same procedure as in steps C1b–g.
8. Following the same procedure as in steps C2–4, perform in-fusion cloning and transform the constructed plasmid vector into E. coli, followed by colony PCR.
To amplify the pegRNA sequence, perform colony PCR using Spacer_OPTN_forward and 3’extension+Linker_OPTN_reverse primers.
9. Following the same procedure as in steps C5–6, culture the transformed E. coli, purify the plasmid vector, and perform Sanger sequencing to confirm the correct sequence of the inserted fragment. Use the pegRNA_sequence_forward primer, which targets the U6 promoter sequence, to perform Sanger sequencing and analyze the pegRNA sequence downstream of the U6 promoter.
Figure 4. Assembling pegRNA using overlap PCR. A. The process of assembling pegRNA using overlap PCR. The homologous sequences of each component anneal to each other, initiating PCR and ultimately synthesizing the pegRNA spanning the entire region from the spacer to the linker sequences. B. Agarose gel electrophoresis image displaying bands corresponding to the PCR-amplified pegRNA components (spacer, spCas9 scaffold, and 3’extension+linker) and the assembled pegRNA. The target-specific band size is around 50 bp in the spacer, 80 bp in the spCas9 scaffold, 60 bp in the 3’extension+linker, and 170 bp in the assembled pegRNA. M: 100 bp DNA molecular weight marker.
D. Identification of highly efficient pegRNAs by Sanger sequencing analysis in HEK293T cells
To enhance the success rate of generating genome-edited cell lines in cell types with low prime editing efficiency, such as iPS cells, it is crucial to select efficient pegRNAs in advance. Therefore, it is recommended to conduct preliminary experiments in cell types known to have high prime editing efficiency, like HEK293T cells, to narrow down efficient pegRNAs.
Specifically, the designed pegRNA and prime editor expression vectors are introduced into HEK293T cells, and the sequence of the target locus is analyzed. Sanger sequencing, which is a cost-effective and simple method, is used for sequence analysis. By comparing the analysis results using wave analysis software [e.g., ICE (Inference of CRISPR Edits) analysis], an approximate estimation of editing efficiency can be made based on the difference in the base sequence from the wild type.
While it is possible to skip this preliminary experiment and start experiments with iPS cells immediately, due to the low editing efficiency of iPS cells, it is recommended to perform more sensitive amplicon sequencing, although it is more costly. Amplicon sequencing involves PCR amplification of the target region followed by next-generation sequencing, allowing for more accurate evaluation of editing efficiency.
1. Ensure the health and viability of HEK293T cell cultures through routine maintenance.
a. Maintain HEK293T cells in DMEM supplemented with 10% FBS and 1% penicillin-streptomycin. Typically, 2 mL of medium is used per well of a 6-well plate. Incubate cells at 37 °C in a humidified atmosphere containing 5% CO2.
Note: Aim for a confluency of 70%–90% with a cell density of approximately 2 × 105–5 × 105 cells/cm2.
b. Observe cells daily under a phase-contrast microscope and change the medium every 2 days.
c. When cells reach 70%–90% confluence, wash the culture plate with 1 mL of DPBS per well of a 6-well plate and treat with 1 mL of 0.05% trypsin/EDTA at 37 °C for approximately 2–3 min. Neutralize the trypsin by adding 2 mL of DMEM (+10% FBS). Subsequently, transfer the cell suspension to a 15 mL centrifuge tube. After centrifuging at 200× g for 3 min, remove the supernatant. Gently resuspend the cells, avoiding bubbles, and subculture at a ratio of 1:100 to 1:50.
2. Co-transfect HEK293T cells with pegRNA and PEmax expression vectors to introduce an SNV. Subsequently, perform puromycin selection to enrich for transfected cells (Figure 5).
Note: To account for potential pipetting errors, prepare 1.2 times the required amount of all transfection reagents.
a. Seed a suspension of viable HEK293T cells at a density of 2.0 × 104 cells per well in a 48-well plate.
b. The following day, when the cells reach 20%–30% confluency, perform co-transfection with pegRNA and PEmax expression vectors. Replace the medium 1 h before transfection.
c. Per each well of a 48-well plate, add 125 ng each of pegRNA and PEmax expression vectors to a sterile 0.5 mL microcentrifuge tube containing 15 μL of DMEM. Set up a negative control with only PEmax expression vector.
d. Per each well of a 48-well plate, add 0.75 μL of PolyJet transfection reagent to a sterile 0.5 mL microcentrifuge tube containing 15 μL of DMEM.
e. Per each well of a 48-well plate, mix 15 μL of plasmid DNA/DMEM and 15 μL of PolyJet/DMEM.
f. Incubate at 20–25 °C for 10 min to allow DNA–polymer complex formation.
Note: Never keep the PolyJet/DNA complex longer than 20 min.
g. Per each well of a 48-well plate, add 30 μL of the DNA/PolyJet mixture and gently swirl the plate to ensure even distribution (day 0).
h. Replace the medium with fresh DMEM (+10% FBS) containing 5 µg/mL puromycin 24 h after transfection to initiate selection (day 1).
i. Change the medium daily. On day 4, switch to a puromycin-free medium.
Note 1: To minimize cell detachment, gently change the medium by tilting the plate and allowing the medium to flow slowly down the wall.
Note 2: Non-transfected cells should begin detaching from the plate bottom and dying 24 h post-puromycin treatment. By day 4, there should be very few viable cells remaining in the negative control wells (see Troubleshooting).
Figure 5. Representative image of HEK293T cells after being transfected and selected with puromycin. HEK293T cells were transfected with pegRNA and PEmax expression vectors on day 0 and cultured in a medium containing 5 μg/mL puromycin from days 1 to 4. The puromycin resistance gene in the pegRNA expression vector allowed for the selection of only the transfected cells by day 4. Scale bar indicates 200 μm.
3. Compare the sequences of the target gene locus between wild-type and prime-edited cells to approximate the editing efficiency of each designed pegRNA (Figure 6).
a. As in steps A3–5, extract genomic DNA from the cells, perform PCR on the target gene locus, and confirm the sequence with Sanger sequencing.
b. Input the spacer sequence into the ICE analysis tool (https://ice.synthego.com/#/) and upload both the control file (unedited sample) and experiment file (edited sample) of the Sanger sequencing waveform data. The editing efficiency of each pegRNA is estimated by referring to the discordance plot and examining the divergence level at the target based on the discordance plot before and after prime editing.
Figure 6. Analysis of bulk Sanger sequencing data from prime-edited HEK293T cells. Three pegRNAs with varying editing efficiencies (#1: high; #2: medium; #3: low) were used to prime edit a target gene in HEK293T cells, followed by bulk Sanger sequencing. Each pegRNA was designed to introduce a G-to-A missense mutation at chr9:129,098,086 of the human GRCh38 genome. A. Bulk Sanger sequencing analysis: the target editing site is indicated by the red box in the chromatogram. The editing efficiency of each pegRNA is estimated by comparing the peak height of the edited adenine base at the target site. A higher peak indicates a higher editing efficiency. B. ICE (inference of CRISPR edits) analysis: the discordance plot generated by the ICE software illustrates the base-by-base alignment between the wild-type (control; orange lines) and edited samples (green lines). It quantifies the degree of mismatch between the edited sample and the reference sequence derived from the control trace file. The degree of mismatch at the target locus after prime editing is indicated by the yellow arrow. The y-axis represents the level of discordance at each base, with higher values indicating greater editing efficiency. The vertical black lines represent the DNA cleavage sites introduced by each unique spacer sequence.
E. Co-transfect human iPS cells with pegRNA and PEmax expression vectors to introduce an SNV
A variety of gene delivery methods have been developed to efficiently and safely introduce genetic material into cells, including physical methods (e.g., electroporation, magnetofection), biological methods (e.g., lentiviral transduction), and chemical methods (e.g., lipid- and polymer-based transfection).
Electroporation, a physical method, creates temporary pores in the cell membrane using an electric field, allowing for direct nucleic acid delivery into cells. Its versatility across different cell types makes it suitable for gene delivery into challenging cells such as iPS cells [16]. However, this method requires specialized equipment and has low throughput.
Chemical methods, on the other hand, use polymers or cationic lipids to form complexes with nucleic acids for cellular delivery. These positively charged delivery reagents interact with negatively charged nucleic acids to form complexes that adhere to the cell membrane through electrostatic interactions. Subsequently, these complexes are internalized into cells via endocytosis or phagocytosis. Compared to physical methods, chemical methods offer several advantages: they do not require specialized equipment, are virus-free, and can be performed using standard cell culture facilities. Additionally, the simplicity of the procedure enables simultaneous transfection under multiple conditions.
In this step, a polymer-based chemical transfection method is used to introduce genes into iPS cells. The concentration of the polymer-based transfection reagent has been optimized to minimize cytotoxicity in fragile iPS cells. Following transfection, non-transfected cells are eliminated by puromycin selection, and cells with the transgene are enriched. Single clones are then obtained by limiting dilution, a technique that ensures each clone originates from a single cell. Finally, clones with the desired gene mutation are identified by genotyping.
1. Ensure the health and undifferentiated state of human iPS cell cultures through routine maintenance.
a. Maintain iPS cells on 0.5 µg/cm2 of iMatrix-511-coated plates in StemFit medium supplemented with 1% penicillin-streptomycin. Typically, 1 mL of StemFit medium is used per well of a 12-well plate. Incubate cells at 37 °C in a humidified atmosphere containing 5% CO2. Observe cells daily under a phase-contrast microscope to monitor for any signs of differentiation, such as changes in colony morphology. Change the medium every 2 days or daily if the cells reach 40% confluence.
Note: Aim for a confluency of 60%–80% with a cell density of approximately 5 × 105–1 × 106 cells/cm2.
b. When the cells reach 60%–80% confluency, wash the culture plate with 1 mL of DPBS per well of a 12-well plate and treat with 500 μL of TrypLE Select at 37 °C for approximately 6–9 min. Observe the cells under a microscope until the cell boundaries become distinct and the cells round up. Then, remove the TrypLE Select and add 1 mL of StemFit medium containing 10 µM Y-27632 per well of a 12-well plate. Gently pipette the medium along the bottom of the well to easily collect the cells without damaging them. Subsequently, transfer the cell suspension to a 1.5 mL tube. After centrifuging at 200× g for 3 min, remove the supernatant. Gently resuspend the cells with 1 mL of StemFit medium (+10 µM Y-27632), avoiding bubbles, and subculture with a cell density of 8.5 × 102–3.5 × 103 cells/cm2. The following day, confirm colony formation and replace the medium with Y-27632-free StemFit medium.
2. Co-transfect iPS cells with pegRNA and PEmax expression vectors to introduce an SNV. Subsequently, perform puromycin selection to enrich for transfected cells (Figure 7).
Note: To account for potential pipetting errors, prepare 1.2 times the required amount of all transfection reagents.
a. When the cells reach 60%–80% confluence and are actively proliferating, perform co-transfection with pegRNA and PEmax expression vectors. Detach the cells from the culture plate using the same procedure as in step E1 and transfer 1× 105 cells to a 0.5 mL tube. Set up cells for a negative control using only the PEmax expression vector. After preparing the transfection reagent as detailed in steps E2b–e, centrifuge the cells at 200× g for 3 min to pellet them.
b. To a sterile 0.5 mL microcentrifuge tube containing 10 μL of DMEM, add 100 ng of pegRNA and 100 ng of PEmax expression vectors per 1 × 105 cells. Set up a negative control with only PEmax expression vector.
c. Per 1 × 105 cells, add 0.8 μL of PolyJet transfection reagent to a sterile 0.5 mL microcentrifuge tube containing 10 μL of DMEM.
d. Per 1 × 105 cells, mix 10 μL of plasmid DNA/DMEM and 10 μL of PolyJet/DMEM.
e. Incubate at 20–25 °C for 10 min to allow DNA–polymer complex formation.
Note: Never keep the PolyJet/DNA complex longer than 20 min. Centrifuge the cells to form a cell pellet immediately before the end of this incubation.
f. Perform a 5-fold dilution of the PolyJet-DNA mixture by adding 20–80 μL of DMEM per 1 × 105 cells.
Note: This is the optimal dilution ratio of the PolyJet–DNA mixture to suppress cytotoxicity in the human iPS cell line 201B7. Since the optimal dilution ratio varies depending on the cell line, it is necessary to optimize the ratio for each cell line as follows: The PolyJet-DNA mixture is prepared according to the manufacturer’s protocol and then serially diluted from the undiluted solution to a 5-fold dilution. Transfection is performed on the iPS cell line at each dilution. The maximum concentration of the transfection reagent that allows for >90% cell viability the following day is determined to be the optimal one.
g. After removing the supernatant, add 100 μL of the diluted PolyJet–DNA mixture per 1 × 105 cells and gently resuspend the cells by pipetting.
h. Incubate the cells at 37 °C for 20 min.
i. Add 10–20 μL of cell suspension per well (approximately 5 × 103–1 × 104 cells) to 12-well plates pre-coated with 0.5 µg/cm2 of iMatrix-511 and containing 1 mL of StemFit medium supplemented with 10 μM Y-27632 (day 0). Pipette gently with a 1 mL pipette to evenly distribute the cells throughout the well.
j. Replace the medium with 1 mL of fresh StemFit medium containing 0.25 μg/mL puromycin and 10 μM Y-27632 24 h after transfection to initiate selection (day 1).
Note: The optimal concentration of puromycin varies depending on the iPS cell line, so it is necessary to optimize it for your specific cell line as follows: Transfect cells with or without a puromycin expression vector, and then treat them with 0.1–0.5 µg/mL of puromycin. The optimal concentration of puromycin is considered to be the concentration at which no viable cells remain in the negative control, but viable colonies are observed in the transfected wells.
k. Change the medium daily. On day 4, switch to a puromycin-free StemFit medium (+10 μM Y-27632).
Note 1: To minimize cell detachment, gently change the medium by tilting the plate and allowing the medium to flow slowly down the wall.
Note 2: Non-transfected cells should begin detaching from the plate bottom and dying 24 h after puromycin treatment. By day 4, there should be very few viable cells remaining in the negative control wells (see Troubleshooting).
l. Change the medium with fresh StemFit medium every 1–2 days until the cell colonies reach a diameter of 500 μm or more (Figure 7).
Note: In each well of the 12-well plate, it is expected that more than 20–30 cell colonies will be formed and that the total number of cells will be more than 2 × 104–4 × 104. However, at this stage, the transfected cells have only been selected, so it is not certain that each colony is a genetically identical single clone.
Figure 7. Representative image of human iPS cells after being transfected and selected with puromycin. The human iPS cell line 201B7 was transfected with pegRNA and PEmax expression vectors on day 0 and cultured in a medium containing 0.25 μg/mL puromycin from days 1 to 4. The puromycin resistance gene in the pegRNA expression vector allowed for the selection of only the transfected cells by day 4. Cell colonies of the transfected and selected cells reached a diameter of approximately 500 µm on days 7–10. Scale bar indicates 500 μm.
3. Detach the cell colonies formed in step E2 and seed them into a 96-well plate by limiting dilution to obtain single clones.
a. Following step E1, trypsinize the cells using TrypLE Select. Prepare a single-cell suspension at a concentration of 10 cells/mL in a reservoir. Mix the cell suspension well to homogenize it using a large pipette and seed 100 μL aliquots into a 96-well plate coated with 0.5 μg/cm2 of iMatrix-511 to achieve a cell density of approximately 1 cell per well.
b. Observe the wells daily and record the wells in which colonies have formed. As a backup, it is also possible to use a part of the cell for expansion culture.
Note 1: If you observe two or more colonies in a single well, or if a colony is significantly larger than others (more than twice the diameter), it may not be a single clone. However, since it is possible that the desired mutation has been introduced, as will be confirmed by the subsequent sequencing analysis, do not exclude these wells but proceed with the following steps. After sequencing analysis, if necessary, perform limiting dilution again to obtain single clones.
Note 2: Ensure that 200 μL of media is added per well to minimize the meniscus effect and allow for clear observation of the edges of the wells. Expect to see colony formation in at least 20%–40% of the wells.
4. Culture until colonies reach a diameter of 400–600 μm. Re-disperse the formed colonies and seed them into 96- or 48-well plates for expansion culturing.
Note: At this step, it is also possible to use half of the cells for direct PCR in section F.
5. When the cells reach 60%–80% confluence, detach the cells and aliquot the cell suspension into 0.5 mL tubes for cell banking and genotyping. For cell banking, cell pellets are resuspended in 200 μL of STEM-CELLBANKER and stored at -80 °C. For genotyping, cell pellets are stored at -30 °C. As a backup, it is also possible to use a part of the cell for expansion culture.
Note 1: For genotyping, a cell pellet equivalent to 1/2 to 1/20 of the cells from a single well of a 96-well plate is sufficient.
Note 2: Ensure that each clone’s cell bank and genotyping frozen pellet are numbered consistently.
F. Genotyping of single clones with direct PCR
Establishing and identifying a specific cell line with the desired genetic mutation from numerous candidate cell lines is a time-consuming and laborious process. This paper introduces an improved method for streamlining this process: rapid detection of gene mutations using direct PCR with cell pellets. This method eliminates the need for the conventional, time-consuming process of genomic DNA purification, allowing for faster identification of the target cell line. Specifically, direct PCR and Sanger sequencing are used to amplify the target gene region and directly confirm the presence or absence of the desired mutation.
1. Perform direct PCR on the frozen cell pellet from step E5 to amplify the target locus for genotyping.
a. Prepare a 30 μL PCR reaction mixture for each sample (Table 10). Prepare a master mix of 1.1 to 1.2 times the total volume of PCR mix required and dispense 24 μL into each PCR tube.
To amplify the region of the OPTN p.(Asn51Thr) missense mutation, perform PCR using OPTN_genotyping_forward and OPTN_genotyping_reverse primers.
Table 10. PCR reaction mixture components
Reagent Final concentration Volume
KOD One PCR master mix (dye-free 2×PCR master mix) 1× 15 μL
Forward primer (2 μM) 0.3 μM 4.5 μL
Reverse primer (2 μM) 0.3 μM 4.5 μL
Sample n/a 1 μL
Nuclease-free water n/a To 30 μL
Total n/a μL
b. Add 10 μL of DPBS to the cell pellet from step E5 and mix well by pipetting or pulse vortexing.
c. Add 1 μL of each cell sample to the PCR reaction mixture, mix by inverting, and then spin down.
d. Perform PCR according to the procedure in steps A1–5, confirm the amplification product by electrophoresis, purify it, and perform Sanger sequencing. By doing so, the genotype of each single-cell line can be identified.
Note: When the PCR product yield is low, increasing the cycle number to approximately 40 can help amplify the product. If this is insufficient, re-PCR using the primary PCR product as a template can be an effective strategy. However, it is important to note that increasing the cycle number also increases the risk of amplification errors, potentially increasing the risk of misinterpreting sequence analysis. Alternatively, although additional steps are required, the efficiency of PCR amplification can be improved by using purified genomic DNA derived from cell pellets as a PCR template.
Figure 8. Direct PCR amplification of the target gene from cell pellets. Agarose gel electrophoresis image displaying bands corresponding to the target gene (OPTN) amplified by direct PCR with cell pellets suspended in DPBS. Samples 1–10 exhibited successful amplification, while samples 11–20 showed insufficient amplification, suggesting the need for additional PCR reactions. The target-specific band size is around 400 bp. M: 100 bp DNA molecular weight marker: lanes 1–20: samples 1–20.
2. Subsequently, expand the cell line with the desired genetic mutation. To enhance the clone purity of the target mutant, serial dilution can be repeated as needed. At least five aliquots of the obtained clones should be prepared and stored in a -80 °C ultra-low temperature freezer for up to 6 months and in liquid nitrogen for long-term storage. This minimizes the reduction in cell viability upon thawing, enabling repeated use in experiments.
Data analysis
Accurate analysis with Sanger sequencing is essential for the selection of efficient pegRNAs and precise identification of clones with target gene mutations. As described in section A, it is crucial to first confirm that PCR and Sanger sequencing can be successfully performed on a wild-type sample at the target locus. The obtained Sanger sequencing data consists of two parts: FASTA sequence data and waveform data. In particular, waveform data are crucial for evaluating the quality of the sequencing reaction. When analyzing waveform data, it is necessary to carefully check whether the waveform is stable and of high quality throughout the sequence, whether there are multiple overlapping peaks, and whether there are any regions with low-quality values. Since the first and last 50 bases of the read tend to have lower quality, the waveform in the middle region should be evaluated more closely. If abnormalities are observed in the waveform data, it may indicate non-specific amplification in the PCR reaction or low quality of the purified DNA.
Validation of protocol
Using this protocol, we introduced the missense mutation OPTN p.(Asn51Thr) into wild-type human iPS cells and obtained the desired mutant clones with an efficiency of 20%. In an independent experiment targeting the same locus, we achieved 16.7% efficiency. Furthermore, we introduced single nucleotide substitutions at two additional loci, chr9: 129,098,086 (G>A) and chr9: 129,101,921 (G>A), obtaining the desired clones with efficiencies of 2.3% and 5.3%, respectively. These results confirm the efficacy of this protocol for introducing SNVs into human iPS cells.
General notes and troubleshooting
General notes
One limitation of this protocol is the difficulty in selecting highly efficient pegRNAs for human iPS cells in silico.
While deep learning based on experimental data enables the in silico prediction of highly efficient pegRNAs in HEK293T cells, this approach has not been widely applied to human iPS cells, and further data are needed. Currently, preliminary experiments using HEK293T cells or amplicon sequencing of edited iPS cells are considered options for pegRNA selection.
Troubleshooting
Problem 1: Low yield or no amplification of the pegRNA sequence in overlap PCR.
Possible cause: The annealing temperature may be too low due to the base composition and length of the overlap regions in each component.
Solution: Extend the length of the overlap region by an additional 5 base pairs or more.
Problem 2: Cells in the negative control are surviving and proliferating during puromycin selection.
Possible cause: The sensitivity to puromycin can vary depending on the cell line and cell density. Cells at higher densities may be less susceptible to puromycin.
Solution: To ensure the effectiveness of puromycin selection, it is crucial to verify its storage conditions and consider the characteristics of the cell line being used. Puromycin should be stored at -20 °C in the dark to maintain its activity. If the cell line exhibits resistance to puromycin, alternative antibiotics like hygromycin B or G418 can be explored. Additionally, optimizing puromycin concentration and cell density can enhance selection efficiency, especially if there are no underlying issues with the reagents or cell line. The optimal puromycin concentration for iPS cells is between 0.1 and 1 µg/mL, while for HEK293T cells, it ranges from 1 to 10 µg/mL.
Problem 3: No surviving cells are observed after puromycin selection.
Possible cause: The transfection efficiency may be too low, or the sensitivity to puromycin may vary depending on cell line and cell density. Cells with low viability may be more susceptible to puromycin.
Solution: Use a reporter expression vector, such as EGFP, to determine the transfection efficiency of your cell line. If no transfected cells are observed, consider increasing the concentration of the transfection reagent or switching to a different transfection reagent. Alternatively, start selection with half the puromycin concentration specified in the protocol. Additionally, gentle pipetting and other handling procedures during passaging can help maintain cell viability and make them less susceptible to puromycin.
Problem 4: The target gene sequence is not amplified by direct PCR.
Possible causes: Low primer specificity or the genomic sequence may be GC-rich or AT-rich, making it difficult to amplify.
Solution: Raise the annealing temperature, redesign the primers for higher specificity, or use a PCR enzyme designed for amplifying GC-rich or AT-rich sequences, such as PrimeStar GXL DNA polymerase (Takara).
Acknowledgments
We thank Mr. Tim Hilts for editing this document. This work was supported in part by funding sources. This work was supported in part by JSPS KAKENHI Grant Numbers 21K19548 (T.N.), 24K22154 (T.N.), and 24K02600 (T.N.).
Competing interests
The authors declare no competing interests.
Ethical considerations
All experiments were approved by the Expert Committee on Genetic Recombination Experiments Security of Tohoku University (permit no. 2020-094). All experiments involving human cells were conducted in accordance with the Declaration of Helsinki (revised 2013).
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Article Information
Publication history
Received: Oct 27, 2024
Accepted: Dec 12, 2024
Available online: Jan 5, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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5,192 | https://bio-protocol.org/en/bpdetail?id=5192&type=0 | # Bio-Protocol Content
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This is an In Press version of the protocol that has not yet been assigned to an issue.
Peer-reviewed
Campylobacter jejuni Biofilm Assessment by NanoLuc Luciferase Assay
TČ Tjaša Čukajne
PŠ Petra Štravs
OS Orhan Sahin
QZ Qijing Zhang
AB Aleš Berlec
AK Anja Klančnik
In Press, Available online: Jan 08, 2025
DOI: 10.21769/BioProtoc.5192 Views: 77
Reviewed by: Emilia Krypotou Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Applied Microbiology and Biotechnology Dec 2024
Abstract
Campylobacter jejuni, a widespread pathogen found in birds and mammals, poses a significant risk for zoonosis worldwide despite its susceptibility to environmental and food-processing stressors. One of its main survival mechanisms is the formation of biofilms that can withstand various food-processing stressors, which is why efficient methods for assessing biofilms are crucial. Existing methods, including the classical culture-based plate counting method, biomass-staining methods (e.g., crystal violet and safranin), DNA-staining methods, those that use metabolic substrates to detect live bacteria (e.g., tetrazolium salts and resazurin), immunofluorescence with flow cytometry or fluorescence microscopy, and PCR-based methods for quantification of bacterial DNA, are diverse but often lack specificity, sensitivity, and suitability. In response to these limitations, we propose an innovative approach using NanoLuc as a reporter protein. The established protocol involves growing biofilms in microtiter plates, washing unattached cells, adding Nano-Glo luciferase substrate, and measuring bioluminescence. The bacterial concentrations in the biofilms are calculated by linear regression based on the calibration curve generated with known cell concentrations. The NanoLuc protein offers a number of advantages, such as its small size, temperature stability, and highly efficient bioluminescence, enabling rapid, non-invasive, and comprehensive assessment of biofilms together with quantification of a wide range of cell states. Although this method is limited to laboratory use due to the involvement of genetically modified organisms, it provides valuable insights into C. jejuni biofilm dynamics that could indirectly help in the development of improved food safety measures.
Key features
• Quantification of C. jejuni using NanoLuc luciferase.
• The assay is linear in the range of 1.9 × 107 to 1.5 × 108 CFU/mL.
• Following biofilm growth, less than 1 h is required for detection.
• The assay requires genetically modified bacterial strains.
Keywords: Campylobacter jejuni NanoLuc Bioluminescence Biofilm assay Viable but nonculturable Food safety
Graphical overview
Campylobacter jejuni biofilm detection using NanoLuc luciferase assay
Background
Campylobacter jejuni, a pathogen found in the gastrointestinal tract of both domestic and wild birds and mammals, is a major cause of zoonosis worldwide [1]. However, it is sensitive to environmental stressors outside its hosts. To this end, C. jejuni forms biofilms as a survival strategy. Biofilms are characterized by a unique bacterial growth state and act as a protective shield that enables bacterial cells to resist certain environmental stressors—such as temperature fluctuations, desiccation, pH changes, and oxidative stress—commonly found on food preparation surfaces, food products, and water reservoirs [2–5]. Bacteria adapt to environmental stressors by, for example, adopting the nonmotile coccoid form and entering a viable but nonculturable state (VBNC) [6–8]. Viable but nonculturable cells cannot divide in vitro but maintain their membrane integrity and metabolic activity [9], which hinders their detection in quality control tests (a critical aspect of food safety) and may explain the persistence of campylobacteriosis in developed countries despite strict hygiene standards [1]. Studies suggest that bacteria in the VBNC state not only remain viable but also have the potential to recover and cause infections [10,11]. To assess biofilms, conventional cultivation methods and colony counts are generally used and usually complemented with biomass estimation using crystal violet staining [12,13]. The metabolic activity of biofilms can be assessed colorimetrically by monitoring the conversion of tetrazolium salt to formazan [14]. Simultaneously, ATP can be measured using BacTiter-Glo or resazurin fluorescence [8,12,13]. Advanced techniques such as transmission electron microscopy, scanning electron microscopy, and immunofluorescence with flow cytometry or fluorescence confocal laser scanning microscopy provide detailed information on the heterogeneity, cell localization, and structure of biofilms [15–17]. Additionally, PCR-based methods are used [12]. However, most of these methods are expensive, unspecific, technically demanding, and unsuitable for cell quantification. Therefore, new methods that can efficiently and rapidly detect C. jejuni cells would be valuable for advancing research related to food safety and possible applications in the food industry. The reporter protein NanoLuc offers several advantages, such as small size (19 kDa), high temperature stability (Tm = 60 °C), activity over a broad pH range (6–8), and the absence of post-translational modifications or disulfide bonds, which ensures uniform distribution within cells. Its most outstanding feature is its highly efficient bioluminescence. The reaction involves the NanoLuc enzyme, which is constitutively expressed, and the Nano-Glo substrate, commonly referred to as luciferin. Upon addition of the substrate in the presence of oxygen, NanoLuc (as luciferase) catalyzes the oxidation of luciferin and generates oxyluciferin. This molecule releases light in its excited energy state when it returns to its ground state [18,19]. No ATP is required for the luminescence reaction, which helps to minimize background luminescence. This reduction in the background signal ensures that the luminescence signal more accurately reflects the metabolic activity of the cells without external interference [20]. NanoLuc has already been successfully used to monitor and quantify Listeria innocua biofilms [21], and we have adapted the protocol to assess C. jejuni growth and biofilm formation. The protocol provides a comprehensive approach to biofilm monitoring and can be used as an important complement or alternative to conventional biofilm monitoring methods.
Materials and reagents
Biological materials
1. C. jejuni 81-176 (GenBank accession number NC_008787.1) transformed with pMW10_nLuc (GenBank accession number OR958835) (Figure 1)
Figure 1. Scheme of the pMW10_nLuc plasmid. KanR, kanamycin resistance gene; porA, promotor; RBS, ribosome-binding site; nLuc, NanoLuc gene; TT, transcription terminator; BamHI and XbaI, restriction sites.
Reagents
1. Nano-Glo Luciferase Assay System (Promega, catalog number: N1120)
2. Phosphate-buffered saline (PBS) (Sigma-Aldrich, catalog number: P3813)
3. Mueller-Hinton broth (MHB) powder (BD Difco, catalog number: 275730)
4. Mueller-Hinton agar (MHA) powder (BD Difco, catalog number: 225250)
5. Karmali Campylobacter selective supplement (Oxoid, catalog number: SR0167E)
6. Ethanol, 99.8%, for HPLC, absolute (Thermo Scientific, catalog number: 445740010)
7. Kanamycin sulfate from Streptomyces kanamyceticus (BioReagent, Sigma-Aldrich, K1377)
Solutions
1. PBS solution (see Recipes)
2. MHB medium/MHA agar (see Recipes)
3. MHA Karmali agar (see Recipes)
Recipes
1. PBS solution
One pouch of PBS
1 L of purified water
Mix thoroughly. Autoclave at 121 °C for 15 min. Avoid overheating.
2. MHB medium/MHA agar
38 g of MHB powder or 21 g of MHA powder
1 L of purified water
Mix thoroughly. Heat with frequent agitation and boil for 1 min to completely dissolve the MHB or MHA powder. Autoclave at 121 °C for 15 min. Avoid overheating. Pour the MHA agar into sterile Petri dishes.
3. MHA Karmali agar
19 g of MHA powder
500 mL of purified water
One vial of Karmali Campylobacter selective supplement
1 mL of sterile distilled water
1 mL of 99.8% ethanol
Aseptically add 2 mL ethanol/sterile distilled water in a 1:1 ratio to the vial containing the Karmali Campylobacter selective supplement and mix gently to ensure that the contents dissolve completely. Add to 500 mL of the prepared MHA agar cooled to 50 °C (see Recipe 2). Mix well and pour into sterile Petri dishes.
Laboratory supplies
1. White flat-bottomed 96-well plates (Thermo Fisher, Microlite, catalog number: 7417)
2. 50 mL centrifuge tube (Corning, Mini Bioreactor, catalog number: 431720)
3. 1.5 mL microcentrifuge tubes (Eppendorf, Safe-Lock, catalog number: EP0030123611)
4. Pre-sterile 50 mL disposable reservoirs (Biotix, catalog number: EP0030123611)
5. Microplate sealing tape (Corning, catalog number: 3345)
6. Pipette tips 0.1–10 μL (Eppendorf, catalog number: 0030000811), 2–200 μL (Eppendorf, catalog number: 0030000889), 50–1,000 μL (Eppendorf, catalog number: 0030000927)
7. Inoculation loops (Golias, catalog number: EZ08)
Equipment
1. Incubator (Thermo Scientific)
2. Orbital shaker (Thermo Scientific)
3. Luminescence plate reader (BMG LabTech, FLUOstar Omega)
4. Pipette set (Eppendorf, catalog number: 05-403-151)
5. Multichannel pipette (Eppendorf, model: Research Plus, catalog number: 2231300048)
6. Biosafety cabinet (Nuaire)
7. Deep freezer (Haier, model: TwinCool)
8. Spectrophotometer (Bio-Rad, model: SmartSpec 3000, catalog number: 4006168)
9. Centrifuge (Thermo Scientific, model: LEGEND X1R, catalog number: 75004261)
Procedure
A. Reviving glycerol stock of C. jejuni
1. Remove a vial containing a stock culture of C. jejuni, previously transformed with the pMW10_nLuc plasmid via triparental mating [22], from -80 °C and place it on ice. For handling C. jejuni, see General note 1.
2. Using a sterile loop, aseptically transfer half a loop of culture to Karmali-selective plates and incubate the plates at 42 °C under microaerophilic conditions (85% N2, 10% CO2, and 5% O2) for 24 h. Microaerophilic conditions are ensured by blowing the gas mixture into an airtight box, which is then placed in the incubator.
3. Using a cotton swab, respread the colonies grown on Karmali agar to MHA agar containing 30 μg/mL of kanamycin. Incubate the plate at 42 °C under microaerophilic conditions for 24 h.
B. Overnight culture for calibration curve preparation
1. Prepare 15 mL of MHB medium with 30 μg/mL of kanamycin in a 50 mL centrifuge tube.
2. Using a sterile inoculation loop, inoculate a few colonies from the MHA agar plate into the medium prepared above. Vortex the tubes.
3. Incubate at 42 °C under microaerophilic conditions (85% N2, 10% CO2, and 5% O2) with gentle shaking (80 rpm) for 16–20 h.
C. Establishing biofilms in microtiter plates
1. Using a sterile loop, take a few colonies from the MHA agar plate and resuspend them in MHB medium until they reach OD600 = 0.1 (approximately 107 CFU/mL, see General note 2).
2. Transfer 100 μL of this suspension into 9.9 mL of MHB medium to achieve a final concentration of approximately 10 CFU/mL. This will serve as the initial culture.
3. Pipette 200 μL of the initial culture to each well, using three separately prepared samples (biological replicates) with each biological replicate in three wells (technical replicates). This results in nine wells per condition. Cover the plates with sterile microplate sealing tape.
4. Incubate the plates under microaerophilic conditions (85% N2, 10% CO2, and 5% O2) for different time periods (e.g., 4, 8, 24, 48, or 72 h).
5. After incubation, carefully rinse the biofilms in the microtiter plates three times with 100 μL of PBS. After the last wash, add 50 μL of PBS to each well. Proceed with bioluminescence measurements as described in section D. Thaw the Nano-Glo luciferase buffer (from Nano-Glo Luciferase Assay System) and gently mix the Nano-Glo luciferase substrate by pipetting. Allow both the reagents and the cultures in the microtiter plates to equilibrate to room temperature before proceeding with the experiment.
D. Calibration curve preparation
1. Adjust the overnight culture (prepared in section B) using sterile MHB medium to obtain an OD600 corresponding to 1.50 × 108 CFU/mL (see General note 2). This will be the bacterial output suspension for the calibration curve.
2. Prepare five distinct dilutions (0.8, 0.6, 0.4, 0.2, 0.1) in 1.5 mL tubes to create a calibration curve using the prepared bacterial output suspension (1.50 × 108 CFU/mL) as the starting concentration (1.0 or 100%). Prepare each dilution by adding the appropriate volumes of bacterial suspension and sterile growth medium, e.g., tube 0.8: mix 800 μL of bacterial suspension with 200 μL of sterile growth medium, etc.
3. Transfer 50 μL of dilutions to the unused wells of the microtiter plate with biofilms in duplicate.
4. Use sterile MHB medium as a blank, which is placed in three wells. The mean value is subtracted from each measurement.
E. Bioluminescence measurement
1. Thaw the Nano-Glo luciferase buffer and gently mix the Nano-Glo luciferase substrate by pipetting. Allow both the reagents and the cultures in the microtiter plates to equilibrate to room temperature before proceeding with the experiment.
2. Prepare the appropriate amount of substrate by mixing one volume (e.g., 100 μL) of Nano-Glo luciferase assay substrate with 50 volumes (e.g., 5 mL) of Nano-Glo luciferase assay buffer.
3. Quickly add 50 μL of substrate to each well containing the culture and wait 3 min. Then, measure bioluminescence using the plate reader (e.g., Omega FLUOstar) (settings: luminescence mode; integration time, 1 s; settle time, 150 ms).
F. Calculating the concentration of bacteria in biofilms
1. Use Excel to plot the calibration curve with relative light units on the y-axis and CFU/mL on the x-axis. A representative calibration curve is shown in Figure 2.
Figure 2. Representative calibration curve of the Campylobacter jejuni biofilm assay. RLU, relative light units; CFU/mL, colony-forming units per mL.
2. Use linear regression (Excel functions: slope and intercept) to calculate biofilm cell concentrations from measured bioluminescence (see General note 3).
Data analysis
Bioluminescence measurements are saved in Excel format, and Excel can be used to analyze the data. The coefficient of determination (R2) of the calibration curve is determined, and data are considered for analysis only if R2 > 0.95. All measurements are repeated in three biological replicates, each of which is repeated in three technical replicates.
Example for the calculation of the biofilm cell concentration based on the measured bioluminescence (RLU): To calculate biofilm cell concentration from RLU, first create a calibration curve using known CFU/mL dilutions (see section F). For example, the curve equation is y = 0.0002x + 36.49 (k = 0.0002; n = 36.49; x = CFU/mL value; y = RLU value). By rearranging the equation, the CFU/mL is calculated from the RLU value of the samples as in Table 1.
Table 1. CFU/mL calculated from the RLU value of the samples
Sample number RLU CFU/mL
1 11,425 5.69 × 107
2 8,054 4.01 × 107
3 4,983 2.47 × 107
Validation of protocol
This protocol was validated in Čukajne et al. [23], DOI: 10.1007/s00253-024-13383-0.
General notes and troubleshooting
General notes
1. Biosafety level 2 (BSL-2) practices must be followed when handling Campylobacter jejuni, which is classified as a Risk Group 2 pathogen. This includes the use of personal protective equipment such as lab coats and gloves, working in Class II biosafety cabinets to prevent exposure to infectious aerosols, and strictly limiting access to the laboratory to trained personnel. Decontamination of biological waste must be ensured prior to disposal, usually by autoclaving [24].
2. The relationship between OD600 and CFU number should be established in advance by plating bacterial suspensions with different OD600 values (serial dilutions) and counting the colonies. Measured OD600 values depend on the spectrophotometer and other experimental conditions used and are therefore not specified. In our settings, for example, OD600 = 1 corresponds to ~1.50 × 108 CFU/mL.
3. In time-course experiments of biofilm growth, each time point (or each plate measured) requires a new calibration curve prepared from the overnight culture. Therefore, an overnight culture should be prepared one day in advance. By including a calibration curve at each time point, we can account for any variations between plates, ensuring accurate measurements despite the high sensitivity of the method.
Acknowledgments
This study was funded by the Slovenian Research Agency (grant numbers J4-3088, J4-4548, J7-4420, P4-0116, and BI-US/22-24-073). This protocol was derived from Čukajne et al. [23], DOI: 10.1007/s00253-024-13383-0. The authors acknowledge Dr. Eva Lasic for editing and reviewing the manuscript.
Competing interests
The authors declare no competing interests.
References
EFSA, ECDC. (2023). The European Union One Health 2022 Zoonoses Report. EFSA J. 21(12): e8442.
Tram, G., Day, C. J. and Korolik, V. (2020). Bridging the Gap: A Role for Campylobacter jejuni Biofilms. Microorganisms. 8(3): 452.
Joshua, G. W. P., Guthrie-Irons, C., Karlyshev, A. V. and Wren, B. W. (2006). Biofilm formation in Campylobacter jejuni. Microbiology. 152(2): 387–396.
Klančnik, A., Vučković, D., Plankl, M., Abram, M. and Smole Možina, S. (2013). In Vivo Modulation of Campylobacter jejuni Virulence in Response to Environmental Stress. Foodborne Pathog Dis. 10(6): 566–572.
Klančnik, A., Vučković, D., Jamnik, P., Abram, M. and Smole Možina, S. (2014). Stress Response and Virulence of Heat-Stressed Campylobacter jejuni. Microbes Environ. 29(4): 338–345.
Tangwatcharin, P., Chanthachum, S., Khopaibool, P. and Griffiths, M. W. (2006). Morphological and Physiological Responses of Campylobacter jejuni to Stress. J Food Prot. 69(11): 2747–2753.
Klančnik, A., Zorman, T., and Smole Možina, S. (2008). Effects of Low Temperature, Starvation, and Oxidative Stress on the Physiology of Campylobacter jejuni Cells. Croatica Chemica Acta. 81(1): 41–46.
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.
Ramamurthy, T., Ghosh, A., Pazhani, G. P. and Shinoda, S. (2014). Current Perspectives on Viable but Non-Culturable (VBNC) Pathogenic Bacteria. Front Public Health. 2: e00103.
Talibart, R., Denis, M., Castillo, A., Cappelier, J. and Ermel, G. (2000). Survival and recovery of viable but noncultivable forms of Campylobacter in aqueous microcosm. Int J Food Microbiol. 55: 263–267.
Baffone, W., Casaroli, A., Citterio, B., Pierfelici, L., Campana, R., Vittoria, E., Guaglianone, E. and Donelli, G. (2006). Campylobacter jejuni loss of culturability in aqueous microcosms and ability to resuscitate in a mouse model. Int J Food Microbiol. 107(1): 83–91.
Klančnik, A., Šikić Pogačar, M., Trošt, K., Tušek Žnidarič, M., Mozetič Vodopivec, B. and Smole Možina, S. (2017). Anti-Campylobacter activity of resveratrol and an extract from waste Pinot noir grape skins and seeds, and resistance of Camp. jejuni planktonic and biofilm cells, mediated via the CmeABC efflux pump. J Appl Microbiol. 122(1): 65–77.
Trošt, K., Klančnik, A., Mozetič Vodopivec, B., Sternad Lemut, M., Jug Novšak, K., Raspor, P. and Smole Možina, S. (2016). Polyphenol, antioxidant and antimicrobial potential of six different white and red wine grape processing leftovers. J Sci Food Agric. 96(14): 4809–4820.
Ma, L., Feng, J., Zhang, J. and Lu, X. (2022). Campylobacter biofilms. Microbiol Res. 264: 127149.
Oh, E., Andrews, K. J. and Jeon, B. (2018). Enhanced Biofilm Formation by Ferrous and Ferric Iron Through Oxidative Stress in Campylobacter jejuni. Front Microbiol. 9: e01204.
Wagle, B. R., Upadhyay, A., Upadhyaya, I., Shrestha, S., Arsi, K., Liyanage, R., Venkitanarayanan, K., Donoghue, D. J. and Donoghue, A. M. (2019). Trans-Cinnamaldehyde, Eugenol and Carvacrol Reduce Campylobacter jejuni Biofilms and Modulate Expression of Select Genes and Proteins. Front Microbiol. 10: e01837.
Ramić, D., Bucar, F., Kunej, U., Dogša, I., Klančnik, A. and Smole Možina, S. (2021). Antibiofilm Potential of Lavandula Preparations against Campylobacter jejuni. Appl Environ Microbiol. 87(19): e01099–21.
Kaskova, Z. M., Tsarkova, A. S. and Yampolsky, I. V. (2016). 1001 lights: luciferins, luciferases, their mechanisms of action and applications in chemical analysis, biology and medicine. Chem Soc Rev. 45(21): 6048–6077.
Promega Coorporation. (2022). Nano-Glo® Luciferase Assay System Technical Manual.
England, C. G., Ehlerding, E. B. and Cai, W. (2016). NanoLuc: A Small Luciferase Is Brightening Up the Field of Bioluminescence. Bioconjugate Chem. 27(5): 1175–1187.
Berlec, A., Janež, N., Sterniša, M., Klančnik, A. and Sabotič, J. (2022). Listeria innocua Biofilm Assay Using NanoLuc Luciferase. Bio Protoc. 12(3): e4308.
Miller, W. G., Bates, A. H., Horn, S. T., Brandl, M. T., Wachtel, M. R. and Mandrell, R. E. (2000). Detection on Surfaces and in Caco-2 Cells of Campylobacter jejuni Cells Transformed with New gfp, yfp, and cfp Marker Plasmids. Appl Environ Microbiol. 66(12): 5426–5436.
Čukajne, T., Štravs, P., Sahin, O., Zhang, Q., Berlec, A. and Klančnik, A. (2024). Holistic monitoring of Campylobacter jejuni biofilms with NanoLuc bioluminescence. Appl Microbiol Biotechnol. 108(1): 546.
Public Health Agency of Canada. (2024). Campylobacter jejuni - Pathogen safety data sheet. Government of Canada. Retrieved November 29, 2024, from https://www.canada.ca/en/public-health/services/laboratory-biosafety-biosecurity/pathogen-safety-data-sheets-risk-assessment/campylobacter-jejuni.html
Article Information
Publication history
Received: Sep 23, 2024
Accepted: Dec 12, 2024
Available online: Jan 8, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
How to cite
Category
Microbiology > Microbial biofilm
Biological Sciences > Microbiology
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5,193 | https://bio-protocol.org/en/bpdetail?id=5193&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
A Bio-protocol resource
This is an In Press version of the protocol that has not yet been assigned to an issue.
Peer-reviewed
A PDMS-based Microfluidic Chip Assembly for Time-Resolved Cryo-EM (TRCEM) Sample Preparation
XF Xiangsong Feng
JF Joachim Frank
In Press, Available online: Jan 08, 2025
DOI: 10.21769/BioProtoc.5193 Views: 406
Reviewed by: Oneil Girish BhalalaNeha NandwaniMunenori Ishibashi
Ask a question
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Cited by
Original Research Article:
The authors used this protocol in Cell Feb 2024
Abstract
Time-resolved cryo-EM (TRCEM) makes it possible to provide structural and kinetic information on a reaction of biomolecules before the equilibrium is reached. Several TRCEM methods have been developed in the past to obtain key insights into the mechanism of action of molecules and molecular machines on the time scale of tens to hundreds of milliseconds, which is unattainable by the normal blotting method. Here, we present our TRCEM setup utilizing a polydimethylsiloxane (PDMS)-based microfluidics chip assembly, comprising three components: a PDMS-based, internally SiO2-coated micromixer, a glass-capillary microreactor, and a PDMS-based microsprayer for depositing the reaction product onto the EM grid. As we have demonstrated in recent experiments, this setup is capable of addressing problems of severe sample adsorption and ineffective mixing of fluids and leads to highly reproducible results in applications to the study of translation. As an example, we used our TRCEM sample preparation method to investigate the molecular mechanism of ribosome recycling mediated by High frequency of lysogenization X (HflX), which demonstrated the efficacy of the TRCEM device and its capability to yield biologically significant, reproducible information. This protocol has the promise to provide structural and kinetic information on pre-equilibrium intermediates in the 10–1,000 ms time range in applications to many other biological systems.
Key features
• Design and fabrication of high-performance splitting-and-recombination-based micromixer and planar microsprayer.
• Protocol for SiO2 coating on the PDMS surface and fabrication of the microfluidic chip assembly.
• Preparation of time-resolved cryo-EM sample in the time range of 10–1,000 ms.
• Data collection on EM grid covered with droplets from the microsprayer.
Keywords: Microfluidics Micro/nanofabrication Single-particle cryo-EM Time-resolved cryo-EM sample preparation Ribosome HflX
Graphical overview
Background
Time-resolved cryo-electron microscopy (TRCEM) [1] is on the way to become a key technique to unlock the dynamics of biomolecular reactions by providing occupancies (i.e., kinetic data) and structural manifestations of intermediate states in the approximate time range of 10–1,000 ms. The setup of a TRCEM experiment must ensure thorough mixing of the reactants, precise control of reaction time, and fast vitrification, such that reaction intermediates can be trapped and subsequently visualized by single-particle cryo-EM.
Over the past two decades, many TRCEM methods have been developed [2–10]. These can be grouped into two main categories: spraying/mixing and mixing/spraying. In the first category, a sprayer or dispenser device deposits a reactant onto an EM grid that is pre-covered with another reactant. Both mixing and reacting occur on the EM grid, which is then rapidly plunged into the cryogen for fast vitrification. In this case, the length of the reaction time is controlled solely by the duration of the plunging. However, the uniformity of the reaction is not ensured as it relies on the efficiency of diffusive mixing on the grid and is affected by uncontrolled processes at the air–water interface. In the second category, a microfluidic chip combining the functions of mixing, reacting, and spraying is used for controlling the reaction and the deposition of the reaction product onto the grid. Here, the plunging time is kept very short to keep the on-grid time to a minimum. Most importantly, mixing and reacting can be separately manipulated, and the reaction time is defined mainly by the residence time of the sample in the microfluidic chip.
For the reasons stated, we have opted for the mixing/spraying microfluidics-based sample preparation method in our development of TRCEM technology. This development was driven by the need to migrate from silicon, in the initial version of the microfluidic chip [8], to plastic polymers as they are more versatile and cost-effective [4,5,10]. However, this migration faced an important obstacle since polymers such as IP-S and IP-Q photoresins and PDMS are intrinsically hydrophobic and prone to the adsorption of biomolecular samples, thereby impairing control over the reactions. Another problem we needed to address—this one shared with silicon-based chips—was ineffective mixing of fluids due to limited micromixer performance in the laminar flow regime.
In our attempt to overcome these problems, we upgraded our TRCEM setup with a PDMS-based microfluidic chip assembly comprising three modules [11]: 1) a PDMS-based splitting-and-recombination-based (SAR) micromixer with 3D self-crossing channels, which is able to efficiently mix the solutions for uniform initiation of a reaction; 2) a polyimide-coated glass capillary tubing, which ensures precise flow rate control of liquids, to serve as the microreactor whose dimensions define the reaction time; 3) a PDMS-based microsprayer (with a design improved from our previous microsprayer [12]) employed to spray out the droplets of the reaction product onto the EM grid. This sprayer has been demonstrated in our work to be efficient for preparing cryo-EM grids with vitreous ice of controllable, highly consistent thickness. The details of the fabrication of these three modules, their integration into a functioning assembly mounted in the TRCEM apparatus, and the testing of the entire setup with biological samples are described here.
We believe that the protocol contains sufficient detail to allow other teams with appropriate scientific and engineering skills to fabricate the microfluidic chips, assemble the entire functioning apparatus, and duplicate the experiments described.
Materials and reagents
Biological materials
Note: Since this protocol is focused on the preparation of TRCEM grids, the purification of biological materials (E. coli 70S ribosome and HflX) will not be explained here. Note that HflX is known as a universally conserved protein for prokaryotic cells, a GTPase that functions as a rescue factor that splits the 70S ribosome into its 30S and 50S subunits in response to heat shock or exposure to antibiotics. For details, the reader is referred to [11] and literature cited therein.
1. E. coli 70S ribosome (1 mM in buffer solution)
2. HflX (5 mM in buffer solution)
Reagents
1. GTP (Invitrogen, catalog number: 18332015)
2. Methylene blue (Sigma-Aldrich, catalog number: 457250)
3. Tris base (Fisher Scientific, catalog number: BP152-1)
4. Ammonium chloride (NH4Cl) (Sigma-Aldrich, catalog number: 09718)
5. Magnesium acetate [Mg(OAc)2] (Sigma-Aldrich, catalog number: M5661)
6. 2-Mercaptoethanol (βME) (Sigma-Aldrich, catalog number: M6250)
Solutions
1. Methylene blue solution (see Recipes)
2. Buffer solution (see Recipes)
Recipes
1. Methylene blue solution (10 mL)
Use within 3 years and store at room temperature.
Reagent Final concentration Quantity or Volume
Methylene blue 4% (w/v) 2.5 mL
DI water n/a 7.5 mL
Total 1% (w/v) 10 mL
2. Buffer solution (10 mL)
Use within one week and store at 4 °C.
Reagent Final concentration Quantity or Volume
Tris-HCl (1 M, pH 8.0) 20 mM 200 μL
NH4Cl (5 M) 100 mM 200 μL
Mg(OAc)2 (2 M) 10 mM 50 μL
βME (14.3 M) 4 mM 2.8 μL
DI water n/a 9.55 mL
Total n/a 10 mL
Note: Add a predetermined amount of Tris base powder to DI water and adjust it to the desired pH value using HCl to obtain the Tris-HCl buffer (1 M, pH 8.0).
Laboratory supplies
1. Silicon wafer with 100 mm diameter (UniversityWafer, Inc. Category: Silicon, ID: 452)
2. Tweezers (Electron Microscopy Sciences, item number: 0508-L5-PO, https://www.dumonttweezers.com/Tweezer/TweezerStyle/42)
3. AB glue (Loctite, epoxy, dries clear, SKU: 1943587, https://www.loctiteproducts.com/products/central-pdp.html/loctite-clear-epoxy/SAP_0201OIL029V3.html)
4. Single-sided tape (Scotch Magic Tape, 0.75 in. × 650 in., https://www.amazon.com/Scotch-Dispensers-Applications-Invisible-Engineered/dp/B0000DH8HQ?ref_=ast_sto_dp)
5. Double-sided tape (Scotch Double Sided Tape, 0.75 in. × 300 in., https://www.amazon.com/Scotch-Double-Dispenser-Standard-237/dp/B0000DH8IT)
6. Aluminum foil (Reynolds, heavy duty, https://www.reynoldsbrands.com/products/aluminum-foil/heavy-duty-foil)
7. Scissors, surgical scissors, sharp blunt (Fine Science Tools, catalog number: 14001-12, https://www.finescience.com/en-US/Products/Scissors/Standard-Scissors/Surgical-Scissors-Sharp-Blunt/14001-12)
8. PDMS hole punchers (Ted Pella, Inc, Harris Uni-Core, Hole 1.0 mm and Hole 0.5 mm)
9. Cutting Mat (ALVIN Drafting, catalog number: HM1218, https://alvindrafting.com/products/hobby-mat-blue-gray)
10. Glass cutter tool set 2–20 mm pencil style oil feed carbide tip (MOARMOR, https://www.amazon.com/gp/product/B07Y1D243H/ref=ppx_yo_dt_b_search_asin_title?ie=UTF8&psc=1)
11. Sandpaper (S&F STEAD & FAST: https://www.amazon.com/STEAD-FAST-Sanding-Discs-Painting/dp/B0BKKNZRRW S&F STEAD & FAST 6 in. Sanding Discs Hook and Loop 24 Pcs, SKU: SA-SC-005)
12. EM grids (TED PELLA, INC., Quantifoil R 0.6/1 holey carbon copper grid, catalog number: 659-300-CU)
13. Tubings [Polymicro Technologies, TSP180350 with Outer diameter (O.D.) 360 μm and inner diameter (I.D.) 180 μm; TSP100170 with O.D. 170 μm and I.D. 100 μm; TSP075150 with O.D. 150 μm and I.D. 75 μm]
14. Pipette tips (BRANDTECH, 0.5–20 μL, colorless, catalog number: 732104, https://shop.brandtech.com/en/pipette-tips-0-5-20-l-pp-colorless.html; BIOTIX, 20 μL racked, sterilized, catalog number: 63300005 https://biotix.com/products/pipette-tips/xtip4-lts-compatible-pipette-tips/20-%CE%BCl-racked-sterilized-2/)
15. Pipette (Eppendorf, 0.1–5 μL)
16. Ethane gas (Airgas, grade level: research purity)
17. Liquid nitrogen (Airgas, gas grade: Industrial)
18. Vitrification dewar (Nanosoft, SKU: 21021005, https://www.nanosoftmaterials.com/product-page/vitrobot-dewar)
19. Tubes (Thermo Scientific, 15 mL Conical Sterile Polypropylene Centrifuge Tubes, model: 339651, https://www.thermofisher.com/order/catalog/product/339651)
20. Blades (https://uat.garveyproducts.com/Product/Index/7097, model: CUT-40475)
21. Photo mask (Fineline Imaging, Inc. https://www.fineline-imaging.com/)
22. 500 g weight (AMERICAN WEIGH SCALES, model: B00SSK3YNO)
23. Polyimide-coated, fused silica capillary tubing (Polymicro Technologies, TSP075150 with O.D. 150 μm and I.D. 75 μm)
24. SU-8 2050 (KAYAKU ADVANCED MATERIALS INC. SU-8 2050 500ML glass bottle with a cap, catalog number: NC0060520, https://www.fishersci.com/shop/products/NC0060520/NC0060520)
25. SU-8 developer (KAYAKU ADVANCED MATERIALS INC. Photoresist developer solution, catalog number: NC9901158, https://www.fishersci.com/shop/products/su-8-developer-4l/NC9901158)
26. Polydimethylsiloxane (PDMS) (Dow Corning, Sylgard 184, catalog number: 4019862)
27. Acetone (Pharmco-Aaper, Midland Scientific, catalog number: 329000000CSGF)
28. Isopropanol (Fisher Chemical, catalog number: BP26184)
29. Acid piranha [a 3:1 mixture of concentrated sulfuric acid (H2SO4) with hydrogen peroxide (H2O2)]; special protection equipment is required for preparing acid piranha [13], and formal training is highly recommended and required for the sake of safety
30. Nitrogen spray guns with 0.80 filter (Cleanroom World, catalog number: TA-NITRO-4-FT, https://cleanroomworld.com/cleanroom-equipment/nitrogen-spray-guns/90-degree-angle-gun-with-hose-assembly-half-inch-tubing); nitrogen is house-generated in Columbia clean room, industrial grade level
31. Polyimide tape (MYJOR, Model number: B07RZYY2T1, https://www.amazon.com/MYJOR-Temperature-Protect-Printer-Professionals/dp/B07RZYY2T1?th=1)
32. PEEK tubing (INDEX HEALTH & SCIENCE, Yellow, 1/16” × 0.007” × 5 ft, Part no.: 1536, https://www.idex-hs.com/store/product-detail/peek_tubing_yellow_1_16_od_x_007_id_x_5ft/1536)
Equipment
1. Optical microscope 1 (Leica, model: LEITZ DM IL) with a digital camera (AmScope, model: MU 1803-HS)
2. Optical microscope 2 (Nikon, model: ECLIPSE ME600L)
3. Optical microscope 3 (Nikon, model: ECLIPSE LV100ND) with a digital camera (Nikon, model: DS-Ri2)
4. Syringe pump (Cole Palmer Instrument Co., catalog number: 78-0200C)
5. Vacuum system (ZENYTM 3.5CFM 1/4HP Pump: https://www.zeny.us/collections/air-vacuum-pump/, model: VP 125+; Stainless Steel SlickVacSeal Vacuum Chamber: slickvacseal.com, Brand: SlickVacSealTM)
6. Glow-discharge machine (Ted Pella Inc., model: 91000S PELCO easiGlowTM Glow Discharge system for Cryo-EM)
7. Plasma processing system for plasma-enhanced chemical vapor deposition (PECVD) (Oxford Instruments Nanotechnology Tools Limited, model: PlasmaPro® NGP80)
8. Time-resolved apparatus for liquid pumping and pneumatic plunging (developed by Dr. Howard White, Eastern Virginia Medical School). More details about the apparatus can be seen in Section E.
9. Hot plate 1 (VWR International, LLC., catalog number: 97042-634)
10. Hot plate 2 (Electronic Micro Systems Ltd., model: 1000 PRECISION HOT PLATE)
11. Hot plate 3 (Electronic Micro Systems Ltd., model: 1000-1 PRECISION HOT PLATE)
12. Spincoating station (ReynoldsTech Fabricators, Inc., programmable FS5.0 spincoating station)
13. Mask aligner (SÜSS MicroTec, model: SUSS MA6)
14. Plasma system (ANATECH USA https://anatechusa.com/, model: Anatech SCE110)
15. Ultrasonic cleaner (Branson Ultrasonics Corp., model: B1510R-DTH)
Software and datasets
1. Nikon NIS-Elements (version 5.21.03, together with Microscope 2, purchased by Columbia University clean room)
2. Fiji/ImageJ (version 2.1.0/1.53c or another version, free and open source)
3. L-Edit (Win32 8.30, license is required); L-Edit is a mask layout editor for Windows-based platforms. For more details about the software, please contact the SIEMENS company: https://eda.sw.sie mens.com/en-US/ic/ic-custom/ams/l-edit-ic/
4. Electron Microscopy Data Bank (EMDB) (https://www.ebi.ac.uk/emdb/) EMD-29681, EMD-29688, EMD-29687, and EMD-29689
Procedure
A. Fabrication of SAR micromixer
Our micromixer is based on the 3D splitting and recombination (SAR) principle. For 3D models of the SAR micromixer and its mixing performance, please see the previous work by Feng et al. [14,15]. The fabrication of this PDMS-based micromixer is as follows:
1. Mask design
For the fabrication of the micromixer, the conventional soft lithography method is employed. Two layers of microchannel structure (Figure 1A) are required to form a 3D micromixer, as illustrated by the geometrical model in Figure 1B. Based on the design of micromixer and numerical calculations [11,14], we designed the mask shown in Figures 1C and 1D using the software L-Edit. The photo mask we designed (Figure 1C) has 62 units and they all share the same channel structure as the one shown in Figure 1D, so after the soft lithographic process, we are able to obtain 62 PDMS copies of the same channel structure. When we rotate one layer to make its channel face the channel of the other layer, we have the two layers shown in Figure 1A.
Figure 1. Design of the photo mask. A. Two layers of microchannel structure, which are made with the same photoresist and are thus identical. B. Geometrical model of the 3D micromixer. C. Photo mask with 62 units. D. One unit of the mask (C) zoomed in, where the inlet with a microfilter was designed to capture big particles.
2. Printing of photo mask
The mask (Figure 2A) was printed in high resolution (10 μm @ 32K DPI) by the company Fineline Imaging, a printing company offering the highest-quality laser photoplot films and film photomasks on the market (https://www.fineline-imaging.com/). By attaching the photomask film to the glass slide (Figure 2B), we have the mask ready for the following UV lithography.
Note: The mask we designed can be seen in the Supplementary information (File S1).
Figure 2. Photo mask printed by high-resolution printer from Fineline Imaging. A. Photo mask. B. Glass slide. C. Photo mask attached to the glass slide.
3. Fabrication of the microchannels on PDMS material
Note: This process should be conducted in the Yellow Room, which is illuminated with yellow light, causing the room to appear yellow; the lamp is designed to filter out harmful UV wavelengths so unwanted exposure of light-sensitive materials such as photoresist is prevented.
a. Substrate preparation: the substrate wafer should be clean and dry to achieve good process reliability. For best results, the silicon wafer is immersed in piranha solution for 20 min followed by a de-ionized (DI) water rinse.
b. SU-8 coating: 1) Dispense 5 mL of SU-8 photoresist on the wafer and slowly spread it on the wafer by manually tilting the wafer. 2) Spin at 500 rpm for 10 s with acceleration of 100 rpm/s. 3) Spin at 4,000 rpm for 30 s with acceleration of 300 rpm/s to achieve a 40 μm-thick SU-8 film coating on the wafer. The spincoating station, as listed in Equipment section, is used in this step.
c. Soft bake: Put SU-8-coated wafer on the hotplate and heat up first to 65 °C for 3 min and then up to 95 °C for 6 min. Then cool down to room temperature.
d. Exposure: Use the photomask (Figure 2C) and apply an exposure dose of 160 mJ/cm2 using the mask aligner (SÜSS MicroTec, SUSS MA6).
e. Post-exposure bake: Put the wafer on the hotplate and heat up to 65 °C for 1 min and then up to 95 °C for 6 min. Then cool down to room temperature.
f. Development: Spray KAYAKU SU-8 developer onto the SU-8 wafer within 5 min, then clean and clear structures coming up.
g. Rinse and dry: Spray and wash the developed image with fresh solution of isopropyl alcohol (IPA) for 10 s. Air-dry in a gentle way with nitrogen gas.
h. Hard-bake (final cure): Bake at 160 °C for 10 min. The SU-8 mold is obtained as shown in Figure 3A.
i. Use aluminum foil to form a container with the SU-8 mold on the bottom. Use the polyimide tape to surround the edge of the wafer and to fix it in place (Figure 3B).
j. Mix 30 g of base and 3 g of curing agent in a cup and pour the mixture (~30.5 g) into the container mentioned above.
h. Put the container into the vacuum for degassing and cure it on the hotplate at 100 °C for 1 h. Then cool down the PDMS.
i. Peel off the cured PDMS replica from the SU-8 mold as shown in Figure 3C.
Note: The SU-8 mold can be reused for a couple of years if it is properly preserved.
j. Follow the boundary of each PDMS unit, as highlighted in Figure 3C, and cut the PDMS replica into pairs of PDMS slabs with the same microchannel structure as shown in Figure 1D. Each pair (Figure 3D) is ready for the micromixer fabrication.
Note: One PDMS replica (Figure 3C) from the SU-8 mold produces 31 pairs of PDMS slabs, which ideally enables us to obtain 31 micromixers.
Figure 3. Fabrication of the microchannels from PDMS material. A. SU-8 mold. B. Container with SU-8 mold on the bottom. C. PDMS replica peeled off from the SU-8 mold, with the boundary of a PDMS unit highlighted. D. Example of seven pairs of PDMS slabs. (Note that the white areas in A, B, and C are from the reflection of the ceiling light, not from patterns on the mold).
4. Alignment and bonding of micromixing channel
a. Clean the surface of the PDMS with tape.
b. Plasma-treat the surfaces of two PDMS slabs with the microchannel side up (15 s Ar at 100 W and 1 min O2 at 100 W).
c. Apply a drop of DI water (4 μL) on the surface of the bottom layer for lubrication.
d. Align the microchannels on the PDMS slabs under the microscope as shown in Figure 4.
Note: A lot of practice is required, and the alignment should be completed within 1 min for best bonding quality.
Figure 4. Alignment during the fabrication of the 3-D PDMS micromixer. A, B. Top and bottom layer of microchannel. C. Misalignment in the beginning. D. Move top layer to align the microchannel with reference to inlets and outlet. E. Well-aligned microchannel.
e. Put the aligned PDMS slabs in vacuum (< 50 mTorr) for 2 min after alignment to remove extra water in between the PDMS slabs; this procedure pre-bonds the PDMS slabs.
f. Check if misalignment has occurred as a result of exposure to the vacuum.
g. Cover the pre-bonded PDMS slabs with small pieces of aluminum foil (Figure 5A).
Figure 5. Fabrication of the micromixer. A. Cover the aligned PDMS slabs with small pieces of aluminum foil. B. Put a 500 g weight on the aligned PDMS slabs during baking. C. Raw tubing (O.D. 150 μm and I.D. 75 μm) with coarse opening. D. Sanded tubing (O.D. 150 μm and I.D. 75 μm); the length depends on the need. E. Application of a bead of glue onto the corner between the tubing and PDMS surface, as marked by the dashed curve. F. The glue covers the corner after application. G. Examples of six micromixers with inlet and outlet tubings inserted and glued. H, J. Two tubings are inserted and connected to the microfilters and well glued. I. One glass capillary tubing is inserted, connected to the outlet, and well glued.
h. Bake pre-bonded PDMS slabs on the hotplate at 120 °C for 10 min with a 500 g weight on it (Figure 5B) to further strengthen the bonding and then check the alignment under the microscope. This is the fully formed micromixer.
i. Apply plasma treatment to the surfaces of the small glass slide and the micromixer (15 s Ar at 100 W and 1 min O2 100 W).
j. Apply a small drop of DI water (2 μL) on the surface of the glass slide for lubrication.
k. Put the bonded PDMS slab onto the surface of the glass slide and move it to its center.
l. Bake at 120 °C for 2 min.
m. Cut the tubing (O.D. 150 μm and I.D. 75 μm) into pieces (Figure 5C) and sand the openings using sandpapers (3,000 grit and then 4,000 grit) such that the ends will become smooth and flat (Figure 5D).
n. Insert well-polished tubings (Figure 5D) into the two inlets and the outlet of the micromixer (Figure 5E). Here, a polyimide-coated, fused silica capillary tubing (O.D. 150 μm; I.D. 75 μm) is used. Note that another type of tubing (O.D. 170 μm; I.D. 100 μm) can also be used for this step, and the length of the tubing depends on the time point desired. Note that the tubings with 2.5, 4.5, and 5 mm in length are used according to our needs.
o. Apply a very tiny amount of AB glue (size of bead: ~1 mm in diameter) on the areas between the tubing and the micromixer. The area is marked by the dashed curve (Figure 5E, F). The glue is filled between the tubing and the PDMS channel, as shown in Figure 5H–J. (After mixing the AB glue, wait for 1 min before applying it to the target area to prevent the glue from flowing too quickly and entering the tubing, thereby blocking it).
5. SiO2 coating
a. Put the PDMS micromixers (as shown in Figures 5C and 6A) into the chamber of the PlasmaPro® NGP80 PECVD machine.
Figure 6. SiO2 coating. A. Examples of four micromixers fabricated following the above-mentioned step A4. B. PECVD machine of model PlasmaPro® NGP80. C. Monitoring of the coating process.
b. Stabilize the stage temperature at 300 °C.
c. Let in the source gasses SiH4 (170 sccm) + N2O (710 sccm) at a vacuum pressure of 200 mTorr.
d. Apply high radio frequency (RF) power (50 W) to create plasma inside the process chamber. Set the coating strike time for 20 min.
e. Vent and open the chamber and store the coated micromixers in a Petri dish in a normal room for future use.
6. Testing of the micromixer
a. Use the IDEX PEEK tubing of 6 mm in length as the connector to connect one end of the polyimide tubings (O.D. 150 μm; I.D. 75 μm; 30 cm in length) to the inlet of the micromixer (Figure 7A); the other end is inserted into another piece of the IDEX PEEK tubing of 4 cm in length, which will be connected to the syringes pump. Apply the AB glue on the junction regions; the process is similar to the step shown in Figure 5E.
Figure 7. Setup for testing the micromixer. A. Fabricated micromixer with tubings for connection to the syringe pump. B. Equipment required for testing, comprising of a syringe pump, a microscope, a camera, and a laptop.
b. Build the setup for testing, which includes a syringe pump and a microscope with a camera and laptop (Figure 7B).
c. Connect the micromixer inlets to the syringes via glass tubings with O.D. 150 μm and I.D. 75 μm.
d. Connect the micromixer outlet to a tube that collects the mixture solution.
e. Introduce the dye solution (or fluorescent solution) and DI water into the micromixer.
f. Run the total flow rates at 2, 4, 6, 8, and 9 μL/s (Figure 8B–F) and capture the mixing status for each setting.
Figure 8. Testing of micromixer. A. Micromixing channel at a flow rate of 0 μL/s. B, C, D, E, and F. Images of mixing statuses at flow rates of 2, 4, 6, 8, and 9 μL/s, respectively. G. Intensity distributions at inlet and outlet when the flow rate is 6 μL/s, as shown in Figure 8D. H, I. Microfilters at the inlets for capturing big particles above 20 μm in diameter.
g. Check if there is evidence of any expansion in the mixing area of microchannels at 9 μL/s (540 μL/min), which is 50% higher than the intended working flow rate (6 μL/s).
h. Check if there is obvious leakage at 9 μL/s.
i. Measure the intensity distribution across the outlet area (marked by dash lines in Figure 8B–F). As one example, based on Figure 8D, we obtained the intensity distributions at inlet and outlet at a total flow rate of 6 μL/s, as shown in Figure 8G. We found that the intensity at the outlet becomes uniformly distributed after mixing. For the calculation of the mixing efficiency, we refer to Feng et al. [14].
j. Run each flow rate (as shown in Figure 8B–F) five times, each time for 4 s, to check if the micromixer is durable enough.
k. Use only those micromixers that pass this high flow-rate quality check. Discard the others.
B. Fabrication of microsprayer
1. Cut the PDMS into small pieces of PDMS slab (length × width × thickness: 6 × 5 × 4 mm) and use PDMS punchers to make holes to accommodate the sprayer outer tube, liquid tube, and gas tubes (Figure 9A).
Notes:
1. The PDMS slab used here is plain, without any microfluidic channel.
2. Some practice will be required for cutting PDMS and punching holes with PDMS hole punchers of 1 mm and 0.5 mm in diameter.
Figure 9. Process of microsprayer fabrication. A. Trimmed PDMS slab with holes for holding the gas inlet tubings, liquid inlet tubing, and microsprayer nozzle. B. Small pieces of glass slide. C. Bonding of the PDMS slab on the glass slide. D. Insertion of gas tubings into the PDMS slab. E. Application of glue to seal the connection area between the gas tubings in the PDMS slab. F. Raw tubing (O.D. 350 μm and I.D. 180 μm) with coarse opening. G. Sanded tubing (O.D. 350 μm and I.D. 180 μm, length: 6 mm, named: tubing 1), which is used for holding the tubing 2. H. Insertion of tubing 1 into the hole of the PDMS slab, with one raw tubing (O.D. 170 μm and I.D. 100 μm) inside tubing 1. This procedure is used to check if tubing 1 is roughly concentric to the PDMS hole. I. Raw tubing (O.D. 170 μm and I.D. 100 μm) with coarse opening. J. Sanded tubing 2 (O.D. 170 μm and I.D. 100 μm, length: 6 mm) serving as the liquid tubing. K. Insertion of tubing 2 into tubing 1. L. Example of seven copies obtained by repeating the steps above. M. Outer tubing 3 of microsprayer with a length of 2 mm. N. Insertion of tubing 3 into the PDMS slab to form the microsprayer nozzle. O. Tubings 3 and 2 are aligned to be concentric, tubing 2 as the inner tube for the liquid, and tubing 3 as the outer tube for the driving gas. P. Example of seven microsprayers obtained by repeating the steps above. Q. Gas tubings. R. Assembled microsprayer ready for use.
2. Cut the normal glass slide (length × width × thickness: 75 × 25 × 1 mm) into small pieces with length × width × thickness: 25 × 7 × 1 mm (Figure 9B) using a glass cutting tool.
Note: Different glass cutter tools can be used, and some practice will be required.
3. Apply plasma treatment to the surface of the PDMS slab (Figure 9A) and small glass slide (Figure 9B) (2 min, 15 mA at air in the EasiGlow cleaning machine).
4. Apply a small drop of DI water (2 μL) on the surface of the small glass slide for lubrication. Align the PDMS slab on the glass slide as illustrated in Figure 9C and then bake the assembly on a hot plate at 100 °C for 2 min for strong bonding.
5. Cut the BRANDTECH pipette tips into a shape as shown in Figure 9D and insert them into the side holes of the PDMS slab. They will serve as gas inlets for the microsprayer. After plasma treatment on it (2 min, 15 mA at air in EasiGlow cleaning machine), apply some glue (size of bead: ~5 mm in diameter) on the junction area of the connection (Figure 9E) and wait for the glue to dry.
6. Cut the tubing (O.D. 350 μm and I.D. 180 μm) into pieces with 6 mm length (Figure 9F) and sand the openings using sandpapers (3,000 and then 4,000 grit) such that the ends will become smooth and flat (Figure 9G). In the following, we refer to the tubing obtained as “tubing 1.”
7. Insert tubing 1 into the hole of the PDMS slab as shown in Figure 9H. Then apply a tiny amount (approximate bead size: ~1 mm in diameter) of glue on the junction area between the PDMS and tubing 1 and wait for the glue to dry.
Note: There is one raw tubing (O.D. 170 μm and I.D. 100 μm) inside tubing 1, which is used to check if tubing 1 is roughly concentric with the hole. It will be removed later.
8. Cut the tubing (O.D. 170 μm and I.D. 100 μm) into pieces with 10 mm length (Figure 9I) and sand the openings using sandpapers (3,000 and then 4,000 grit) such that the ends will become smooth and flat (Figure 9J). In the following, we refer to the tubing obtained as “tubing 2.”
9. Insert tubing 2 into tubing 1 and move tubing 2 until 1 mm of it protrudes outside PDMS on the nozzle side. Then apply a tiny amount (size of bead: ~3 mm in diameter) of glue on the junction area between tubing 1 and tubing 2 (Figure 9K) and wait for the glue to dry. Figure 9L shows several copies obtained by repeating the steps above.
10. Cut the BRANDTECH pipette tips into a shape with 2 mm length, as shown in Figure 9M, to be used as the microsprayer outer tube. In the following, we refer to the tubing obtained as “tubing 3.”
11. Insert tubing 3 into the hole of the PDMS slab (Figure 9N) and carefully align tubing 3 with tubing 2 under the microscope to make them concentric, as shown in Figure 9O.
Note: Tubings 2 and 3 form the microsprayer, with tubing 2 for the liquid and tubing 3 for the driving gas.
12. Following step B11, perform the plasma treatment on the sprayer assembly (2 min 15 mA at air in EasiGlow cleaning machine), apply some glue (bead size: 1 mm in diameter) to the junction area between tubing 3 and the rim of the PDMS hole, and wait for the glue to dry. Figure 9P shows several copies obtained by repeating the steps above.
13. Cut the soft tubing (O.D. 2.4 mm and I.D. 0.8 mm) into pieces with 15 cm length. Cut the BRANDTECH pipette tip with a length of 8 mm and bend them to be in a curved shape. Assemble the soft tubing and the curved tip as shown in Figure 9Q. In the following, we refer to the tubing obtained as “gas tubing.”
14. Insert the two gas tubings into the two gas inlets of the microsprayer (Figure 9R) and apply some glue (bead size: ~4 mm in diameter) to the junction area between the gas tubings and gas inlets. Wait for the glue to dry.
15. Now, the microsprayer is ready to be used in the fabrication of the entire chip assembly with a time point longer than 20 ms.
C. Fabrication of the microfluidic chip assembly
C1. For time points longer than 20 ms
1. Choose proper glass capillary tubing to connect the micromixer with the microsprayer. Here, for demonstration, a tubing with O.D. 350 μm and I.D. 180 μm is used and cut to a length of 83 mm; we will call it the microreactor tubing. (Note: Since the reaction time is related to the volume of the microreactor tubing, the length and diameter of the microcapillary tubing can be changed to achieve different time points. The micromixer outlet tubing and microsprayer inlet tubing must be changed in size for proper connection.)
2. Insert the outlet tubing of the micromixer (O.D. 170 and I.D. 100 μm) into the microreactor tubing (O.D. 350 μm and I.D. 180 μm) on one end and insert the liquid tubing of the microsprayer into the microreactor tubing (O.D. 350 μm and I.D. 180 μm) on the other end. The micromixer, the microreactor, and the microsprayer are now connected to each other in the complete assembly (Figure 10).
Figure 10. Fabrication of microfluidic chip assembly with time points longer than 20 ms
3. Apply a tiny amount (bead size: ~1 mm in diameter) of glue to the connection area between the micromixer and the microreactor and to the area between the microreactor and microsprayer, and then wait for the glue to dry.
Note: After mixing the AB glue, wait for 1 min to reduce the fluidity of the glue before applying it to the areas of interest, to prevent the glue from clogging the microreactor tubing. Regarding the timing for applying the glue, first do some testing on separate raw tubings with the same diameters as specified in steps 1 and 2 to check how the glue flows inside the gap between the tubings and when it stops flowing.
4. Estimate and calculate the reaction time based on the design of the fabricated chip. As we stated in our work [11], the total reaction time is comprised of the mixing time in the micromixer, the reaction time in the microcapillary reactor, the spraying time, the plunging time, and the reaction time during the vitrification process. In this example (Figure 10), a total reaction time of 350 ms will be obtained.
Notes:
1. The volume taken up by the inserted tubing on the connection region needs to be subtracted.
2. In our work [11], following this process, we fabricated a group of chips with three time points: 25 ms, 140 ms, and 900 ms.
5. Check the sample adsorption with the E. coli 70S ribosome by using the setup as depicted in Figure 13. Collect the sample before and after passing the device and measure the concentration using a Nanodrop UV-Vis spectrophotometer. In this testing, 94% of the initial concentration can be retained using the SiO2-coated chip. (Gas pressure for the spray is 8 psi and the total liquid flowrate is 6 μL/s; for more details on different coatings, see [11].)
C2. For time points shorter than 20 ms
1. Follow the steps shown in Figure 9A to prepare the PDMS slab with holes (one front hole, two side holes, and one back hole), and trim it to have a smaller size (length × width × thickness: 5 × 4 × 2 mm). Here, we call this PDMS slab a nozzle PDMS slab. The front hole is used for accommodating the sprayer outer tube, the back hole is for the liquid tube, and the side holes are for gas tubes (Figure 11A).
Figure 11. Fabrication of microfluidic chip assembly with time points shorter than 20 ms. A. Prepare a smaller nozzle PDMS slab (5 mm × 4 mm × 2 mm) with holes for holding the gas inlet tubings, liquid inlet tubing, and microsprayer nozzle. B. Insert gas tubes into the PDMS slab. C. Insert outlet liquid tubing into the back hole of the nozzle PDMS slab. D. Fix the micromixer on the side of a table to ensure that the front hole of the nozzle PDMS slab faces up. E. Apply glue to seal the contact area between the micromixer and the nozzle PDMS slab. F. The nozzle PDMS is glued together with the micromixer. G. Outer tube with a length of 2.5 mm for the microsprayer. H. Insert the outer tube into the front hole of the nozzle PDMS slab to form the microsprayer. I. Outer tube and inner tube (i.e., outlet liquid tubing of the micromixer) are aligned to be concentric. J. Fix the micromixer on the side of a table to ensure that the outer tube of the microsprayer faces up and apply the AB glue to seal the connection between the outer tube and the nozzle PDMS slab. K. The microsprayer nozzle with AB glue on the corner between the outer tube and the nozzle PDMS slab. L. Put PEEK tubings on the inlets of the micromixer. M. Gas tubings. N. Insert the gas tubings into the gas tubes. O. Insert the liquid tubings into the connector PEEK tubings. P. Application of the AB glue to the dash circled areas. Q. Assembled microfluidic chip assembly with time points shorter than 20 ms, ready for use.
2. Cut the BRANDTECH pipette tip with a length of 8 mm and bend into a curved shape. Then insert the tip into each side hole of the nozzle PDMS slab as shown in Figure 11B.
3. Take one micromixer with a liquid tube as its outlet (Figure 5G) and insert the liquid tube into the back hole of the nozzle PDMS slab. Then, the two surfaces of the nozzle PDMS and micromixer PDMS slab can make contact with each other as shown in Figure 11C.
4. Apply plasma treatment to the assembly of the micromixer and nozzle PDMS slab (Figure 11C) (2 min, 15 mA at air in EasiGlow cleaning machine). Then fix the micromixer on the side of the table to ensure that the front hole of the nozzle PDMS slab faces up (Figure 11D) and apply some AB glue (bead size: 1 mm in diameter) a few times until the AB glue will cover the corner between the nozzle PDMS and micromixer PDMS slab, as dash-circled in Figure 11E. After the glue becomes dry, the nozzle PDMS is attached to the micromixer (Figure 11F).
5. Cut the BRANDTECH pipette tips into a shape with 2.5 mm length, as shown in Figure 11G, to be used as the microsprayer outer tube.
6. Insert the outer tube into the front hole of the PDMS slab (Figure 11H) and carefully align the outer tube with the liquid tube under the microscope to make them concentric, as shown in Figure 11I.
7. Fix the micromixer on the side of a table to ensure that the outer tube of the microsprayer faces up (Figure 11J) and apply some AB glue (bead size: 1 mm in diameter) a few times until the AB glue covers the corner between the outer tube and the nozzle PDMS slab (Figure 11K). After the glue becomes dry, the outer tube is attached to the nozzle PDMS slab, and a microsprayer is formed.
8. Cut the IDEX PEEK tubing (O.D. 1/16”, I.D. 0.007”) into pieces with 6 mm length, which serve as the liquid tubing connector, and put two pieces on the inlets of the micromixer as shown in Figure 11L.
9. Cut the soft tubing (O.D. 2.4 mm and I.D. 0.8 mm) into pieces with 15 cm length. Cut the BIOTIX pipette tip with a length of 5 mm. Assemble the soft tubing and the tip (as shown in Figure 11M) to serve as gas tubing.
10. Insert the two gas tubings into the two gas inlets of the microsprayer (Figure 11N).
11. Use tubings (O.D. 150 μm; I.D. 75 μm) with proper length (30 cm) as liquid tubing and insert one end into liquid tubing connectors for connection to inlets of the micromixer (Figure 11O).
12. Apply some glue (bead size: ~ 4 mm in diameter) to the junction area between the gas tubings and gas inlets, to the junction area between the micromixer and liquid tubing connectors, and to the junction area between the liquid tubing connectors and liquid tubing, as dash-circled in Figure 11P. Wait for the glue to dry.
13. Insert the other end of the liquid tubing into a piece of the IDEX PEEK tubing of 4 cm in length (Figure 11R), which will be used to be connected to the syringes pump, and apply the AB glue on the junction regions. The process is similar to the steps shown in Figure 5E.
14. At last, we now have the microfluidic chip assembly with time points shorter than 20 ms (Figure 12).
Note: In our work [11], following this process, we fabricated the chip assembly with a time point of 10 ms.
Figure 12. Entire view of the fabricated microfluidic chip assembly with time points shorter than 20 ms
D. Estimation of the reaction time obtained using this protocol
Based on our previous study [11], the total reaction time can be estimated as t = tm + tr + tf + tp + tv, where tm is the mixing time in the micromixer, tr is the reaction time in the microcapillary reactor, tf is the flight time in the spray, tp is the plunging time, and tv is the reaction time during the vitrification process. Briefly, each part is described as follows:
1. tm is estimated by the equation tm = Vm/Uf, where Vm is the volume of micromixer, and Uf is the total flow rate of the solutions introduced into the micromixer. Here, Vm is 2.80 nL for this micromixer, and Uf is 6 μL/s, so the mixing time is estimated to be ~0.47 ms.
2. tr is estimated by the equation tr = Lr/V, where Lr is the length of the microcapillary reactor, and V is the mean velocity of the fluid.
3. tf is estimated by the equation tf = Df/Vd, where Vd is the averaged velocity of the droplets, and Df is the distance from the microsprayer orifice to the EM grid, fixed at 3.5 mm for our implementation. When the liquid flow rate is 6 μL/s and gas pressure is around 8 psi, Vd is around 6.4 m/s [11], so we obtain a mean droplet flying time of ~0.55 ms.
4. tp is estimated by the equation tp = Hp/Vp, where Vp is the plunging speed, and Hp is the plunging height from the microsprayer orifice to the liquid ethane surface. In this protocol, Vp = 1.9 m/s at N2 gas pressure of 40 psi, and Hp is 10 mm or 15 mm. At Hp of 10 mm, tp is ~5.26 ms; at Hp of 15 mm, tp is ~7.89 ms.
5. tv is estimated by the equation tv = (Troom - 273.15)/CCR, where Troom is the room temperature, and CCR is the critical cooling rate, which is ~105 K/s. So, we can estimate the reaction time in the vitrification process, which amounts to ~0.23 ms.
For this protocol, four chip assemblies are fabricated; all key materials and parameters of the chip assemblies are listed in Table 1. Based on the above-mentioned equation, the reaction times using our chip assemblies could be achieved at 10, 25, 141, and 899 ms (to simplify, we round them to 10, 25, 140, and 900 ms).
Table 1. Materials and parameters for the fabrication of the four chips used in this TR study
Microfluidic chips Capillary tubing (as microreactor) inner diameter (I.D.) and length (μm/mm) Tubing (between micromixer and microreactor) I.D. and length (μm/mm) Tubing (between microreactor and sprayer) I.D. and length (μm/mm) Sprayer–grid distance (mm) Sprayer–cryogen distance (mm) Plunging velocity (m/s)
Chip 1 (10 ms) 75/5.0 N/A N/A 3.5 10 1.9
Chip 2 (25 ms) 200/2.0 75/2.8 75/5.0 3.5 15 1.9
Chip 3 (140 ms) 200/24.0 75/3.0 75/5.4 3.5 15 1.9
Chip 4 (900 ms) 300/75.0 75/2.6 75/5.5 3.5 15 1.9
Note: This table was reported as table S1 in our previous study [11].
E. Preparation of sample for TRCEM
Before the preparation of the sample for TRCEM, we would highly recommend doing non-EM kinetic experiments, which will save much effort and resources. For this study, as detailed in our paper [11], we chose different reaction time points according to our peer researchers’ non-EM kinetic results [16], where they followed the kinetics of ribosome dissociation with HflX in stopped-flow by using light scattering as a reporter. In recent years, single-molecule FRET (smFRET) has been used to capture the dynamic process of a reaction system by measuring distances between fluorescence tags within single molecules even at a nanometer scale [17], and results from such experiments can guide the choice of time points in the TR experiment.
The entire setup for time-resolved grid preparation, based on the computer-operated liquid-pumping and grid plunging apparatus developed by Howard White (Eastern Virginia Medical School) [18], is depicted in Figure 13. As the key component of the TRCEM apparatus, the microfluidic chip assembly is mounted next to the pneumatic plunger (Figure 13), both placed in an environmental chamber [19] that maintains temperature and humidity. The plunger, which is pneumatically driven and controlled, holds the tweezers on which the EM grid is mounted for fast plunging into liquid ethane after passing the spray cone. In addition, the apparatus contains the pumping system for introducing the solutions into the micromixer and the nitrogen gas into the gas inlets of the microsprayer. Finally, it also includes a computer for controlling both the pumping system and the plunger.
Before each spraying experiment, the tweezers are adjusted in a way that the grid held at their tip passes the nozzle and then plunges into the cryogen cup. In all our experiments, the condition inside the chamber was maintained at 80%–90% in relative humidity by a humidifier connected to the chamber and kept at a temperature in the range of 24–26 °C. Compressed nitrogen gas, humidified by passing through two consecutive water tanks, is fed into the microsprayer at a manually regulated gas pressure. Once the gas flow is stable, the solutions are injected into the microfluidic chip assembly by syringe pumps under computer control, and the total liquid flow rate is set at 6 μL/s. At this point, the sprayer starts spraying. Lastly, the EM grid passes through the spray cone and is plunged into liquid ethane. In detail, the procedure of the experiment is as follows:
1. Open the environmental chamber and put the microfluidic chip assembly onto the supporting platform.
2. Mount the tweezers on the plunger and move the plunger up and down to check if the tip of the tweezers will pass through in close vicinity of the opening of the microsprayer nozzle.
3. Align the sprayer nozzle with the tip of the tweezers and move the chip assembly forward and backward (i.e., left and right inside the environmental chamber, Figure 13) to adjust the distance between the sprayer nozzle and the tip of the tweezers to 3.5 mm. After alignment, fix the position of the microfluidic chip using pieces of tape (FisherbrandTM Labeling Tape).
4. Connect the chip assembly to syringes 1 and 2 with the tubing (O.D. 150 μm; I.D. 75 μm) for feeding the sample solution (Figure 13).
Note: The narrow tubing (O.D. 150 μm; I.D. 75 μm) is used to reduce the dead volume of the sample.
5. Connect the chip assembly with the nitrogen gas tank using the tubing (O.D. 2.4 mm and I.D. 0.8 mm) for feeding the gas. There is a gas valve between the gas tank and the microsprayer.
6. Load DI water into the syringes. (As shown in Figure 13, ports A and B for each syringe valve can be used for withdrawing and dispensing liquid, respectively).
Figure 13. Entire setup with the mounted microfluidic device.
Note: More information can also be seen in Figure S1 and Video S1 in our previous paper [11].
7. Run the syringe pumps for three rounds of DI to clean the system. Each time, the total volume to be dispensed is set at 20 μL.
8. Turn on the N2 gas and, at the same time, run the syringe pumps with DI water. Each time, the total volume to be dispensed is set at 20 μL. Increase the gas pressure from 4 to 16 psi and check if the microsprayer keeps working in good condition as shown in Figure 14.
Notes:
1. Normally, we use 8 or 12 psi as working gas pressure.
2. A laser pointer pen is used to check the spray cone.
Figure 14. Testing of the entire setup with the microfluidic device. When increasing the gas pressure from 4 to 16 psi (from A to D), good spray performance can be found at 8 and 12 psi. (Note that a laser pointer pen is used to check the spray cone.)
9. Discharge the DI water from the syringes, load buffer solution into them, and repeat steps E7 and E8. Check whether the microsprayer works using buffer solution.
Note: Check each connection area on the microfluidic chip assembly for possible leakage. It is wise in the planning of an experiment to have one extra chip for backup.
10. Turn on the N2 gas tank, which is connected to the plunger. We set the gas pressure for the operation of the plunging at 40 psi.
11. Observe and check if the plunger is driven by the N2 gas by turning the plunger valve on and off three times using the laptop computer interface.
12. Discharge the buffer solution from the syringes and load two reactant solutions into the empty syringes.
13. Clean the EM-grid with the EasiGlow machine to make the grid hydrophilic (15 mA, 35 s, vacuum pressure: 0.26 m Torr).
14. Hold the EM grid by the tweezers.
15. Mount the tweezers on the plunger.
16. Align the EM grid with the sprayer nozzle and adjust the distance in horizontal direction from the nozzle to the EM grid.
17. Turn on the humidifier to increase the humidity inside the environmental chamber and wait for the humidity to stabilize at around 90%.
18. Turn on the N2 gas for the plunger and the N2 gas for the microsprayer.
19. Move the container with the liquid ethane into the correct position beneath the sprayer nozzle.
20. Turn on the syringe pump and activate the microsprayer to spray. After waiting for the spray plume to stabilize, activate the plunger by the computer, which causes the EM grid to pass through the spray cone and rapidly immerse into the liquid ethane for fast vitrification.
Note: In our experiments, we wait 3 s to allow stabilization of the spray.
21. Turn off the nitrogen gasses and the humidifier.
22. Unmount the tweezers from the plunger and move the EM grid into a grid box in liquid nitrogen.
Note: When transferring the grid from liquid ethane to liquid nitrogen, the faster, the better. As in the blotting method, a lot of practice is needed for mastering fast transfer.
23. Repeat steps E13–E22 to prepare additional grids using the same microfluidic chip assembly. We normally prepare four grids for use with one chip at a specific reaction time point.
24. Unmount the microfluidic chip assembly and mount a new one with a different reaction time point. Then follow the same procedure described above.
Notes:
1. Both prior to and subsequent to a TR experiment, the microfluidic chip assembly needs to be pumped with DI water for cleaning.
2. Buffer solution is always pre-sprayed to ensure the sprayer chip is in a good working state.
3. Each microfluidic chip assembly is used for one given biological reaction only to avoid cross contamination.
Data analysis
The data analysis method we employ already appears in sufficient detail in our published paper [11], so a brief summary suffices here. Four reactions of 10, 25, 140, and 900 ms were prepared via the TRCEM experimental procedure, and 3452, 3598, 3530, and 3603 good micrographs were collected, respectively, using a commercial high-resolution transmission electron microscope. We always pool together all micrographs from an entire series of TRCEM experiments for data processing (see [3]). After correction of beam-induced motion using MotionCor2 [20] and estimation of the contrast transfer function (CTF) for each micrograph using CTFFIND4 [21] in Relion-4.0 [22], particle picking was performed using Topaz [23]. The entire dataset for the four time points contained around 1 million “good” particles. After further rounds of 2D classification, 802,562 particles were selected, which were used to generate a 3D initial model for 3D auto-refinement in Relion-4.0. CTF refinements were done to correct for magnification anisotropy, fourth-order aberrations, per-particle defocus, and per-particle astigmatism, followed by another 3D auto-refinement. Then, 3D classification was performed on the entire pooled dataset without re-alignment, using the angular information from the previous 3D auto-refinement. Seven different classes were found: (1) rotated 70S lacking HflX (r70SnoHflX, 56,894 particles); (2) non-rotated 70S ribosomes lacking HflX (nr70SnoHflX, 96,898 particles); (3) 70S-like intermediate-I bound with HflX (i70SHflX-I, 140,682 particles); (4) 70S-like intermediate-II bound with HflX (i70SHflX-II, 138,296 particles); (5) 70S-like intermediate-III bound with HflX (i70SHflX-III, 113,038 particles); (6) 50S subunit bound with HflX (50SHflX, 62,558 particles); and (7) 30S subunit (58,952 particles) with a total of 667,318 particles. For this pooled dataset, a mixture of particle populations with a reaction time range of 10–900 ms co-exists in each class, and each picked particle was labeled by the time point of the micrograph it originated from, so percentages of particles of different time points contributing to each class can be calculated with respect to the total of 667,318. Detailed description of the analysis can be found in the section “Cryo-EM data processing” and Figure S4D in our previous work [11].
Validation of protocol
This protocol or parts of it has been used and validated in the following research article:
• Bhattacharjee et al. [11]. Time resolution in cryo-EM using a PDMS-based microfluidic chip assembly and its application to the study of HflX-mediated ribosome recycling. Cell (Figure 1; Figure 2; Figures S1, S2; Figure 3 panel M).
We stored the prepared grid in a liquid nitrogen dewar for future imaging. In this study, after the screening check, we obtained around 2 grids on average for each time point. For data collection, we used 3, 1, 2, 1, and 1 grid(s) for 10 ms, 25 ms, 140 ms, 900 ms, and control, respectively.
All the data were collected using a 300-kV Titan Krios (Thermo Fisher Scientific, Waltham, MA) equipped with a K3 direct detector camera (Gatan, Pleasanton, CA). In contrast to normal blotted grids, the droplet-covered TRCEM grids require manual hole and exposure targeting to pick up collectible droplets and avoid thick-ice areas based on the intensity in the hole images, as shown in the left and middle column panels of Figure 15. Some example high-resolution micrographs from TR grids are shown in the right-column panels of Figure 15. All other details about data collection can be found in the article referenced above [11]. As a result, we were able to collect 3,452, 3,598, 3,530, and 3,603 high-resolution micrographs for 10, 25, 140, and 900 ms, respectively, sufficient for investigating the time course of HflX-mediated ribosome recycling.
Figure 15. Data collection. The left column panels are micrographs for hole targeting; the middle column panels are micrographs for exposure targeting; the left column panels are the desired high-resolution micrographs.
As a result of data processing, we found that the HflX, in the presence of GTP, interacts with the 70S ribosome and recycles it by splitting it into its 30S and 50S subunits. Three intermediates (Figure 16A–C) whose occupancies are short-lived yet stretch over two or three time points (Figure 16D) show a stepwise clam-like opening of the 70S ribosome, which allowed us to infer the temporal sequence in which the intersubunit bridges are disrupted. As the density maps were resolved at resolutions around 3 Å, we could build atomic models, allowing the detailed molecular action mechanism of HflX to be inferred. The results demonstrate that this protocol is well adapted to resolve a biological reaction process in the stated time range of 10–1,000 ms. For more information about the biological story, we refer to the original article [11].
Figure 16. Validation of the protocol. A. Initial opening of the 70S ribosome from i70SnoHflX (blue) to i70SHflX-I (brown), forced by the interaction with HflX. B, C. Motion of the 30S subunit from i70SHflX-I (brown) to i70SHflX-II and to i70SHflX-III (yellow), as the 70S ribosome opens stepwise by the action of HflX. The maps of i70SnoHflX, i70SHflX-I, i70SHflX-II, and i70SHflX-III are deposited in the public database as EMD-29681, EMD-29688, EMD-29687, and EMD-29689, respectively. In (A) through (C), all reconstructions are aligned on the 50S subunit. D. Kinetics of the splitting reaction in terms of the number of particles per class (or occupancy of the corresponding state) as a function of time, obtained by 3D classification. Figure 16D was reported as Figure 2M in [11].
General notes and troubleshooting
General notes
1. Over the course of three months, we tested the fabricated microfluidic chip assembly and found that it can be reused multiple times to prepare TRCEM grids if it is properly cleaned and preserved after each experiment.
2. During microfabrication, the volume of the PDMS AB mixture used for curing may vary, and thus the thickness of the PDMS slab for the sprayer may vary from case to case. Therefore, the distance between the sprayer nozzle and the ethane cup needs to be measured for each new chip assembly to estimate the plunging time.
3. Copper Quantifoil EM grids are preferred as they withstand the force of the spray while gold grids tend to be bent.
Troubleshooting
Problem 1: Compared with the conventional blotting method, the TRCEM method requires a substantially bigger volume of sample (10 μL vs. 3 μL) for each grid.
Possible cause: (1) “dead volume”: there is a minimum quantity of fluid required to reside in the whole microfluidic system for ensuring a stabilized spray; (2) much of the spray is wasted in the present setup with a single EM grid as the target; (3) deposition of the sample from the spray plume onto the grid is inefficient, and a lot of the material is wasted.
Solution: While the dead volume problem is difficult to overcome, the waste of active spray can be mitigated by the development of a plunger with multiple pairs of tweezers or a specially designed tweezer manifold to hold several grids at once such that they pass successively through the plume.
Problem 2: Compared with the conventional blotting method, the TRCEM method requires a larger volume of sample (10 μL) for each grid.
Possible cause: The tubings to connect the chip and pump syringe need to be filled with the sample and, compared with the conventional blotting method, a larger volume of sample is required to stabilize the spray before plunging.
Solution: It is essential to develop a plunger with multiple pairs of tweezers or a specially designed tweezer manifold to hold several grids at once.
Problem 3: Data collection on droplet-covered TRCEM grids still relies on manual selection, which is time-consuming.
Possible cause: The droplets have variable sizes and are randomly and often sparsely distributed on the grid surface, and ice thickness can vary from one droplet to another or even within a droplet (see also Problem 4).
Solution: Deep learning–based programs may be able to enhance the effectiveness of the data collection on droplet-sprayed EM grids.
Problem 4: Collectible regions of droplets for high-quality images are limited to the narrow regions that touch the grid bar, which limits the amount of data one can obtain from a single EM grid.
Possible cause: The fast plunging and surface tension of the liquid influence the spreading of the droplet on the EM grid.
Solution: Some specialized EM grids, such as self-wicking or nanowire grids [24] or ultra-flat graphene EM grids [25] are possible options to increase the areas with suitable ice thickness, but these grids need to be strong enough to withstand the force of the gas-assisted spray.
Supplementary information
The following supporting information can be downloaded here:
1. File S1. Mask design
Acknowledgments
This work was supported by a grant from the National Institutes of Health R35GM139453 (to J.F.). All data were collected at the Columbia University Cryo-Electron Microscopy Center (CEC). The microfluidic chips with SiO2 coating were fabricated in the nanofabrication clean room facility of Columbia University. We also thank Swastik De for his helpful discussion. Importantly, the protocol described here is a more detailed account of the experimental setup and experimental results previously published in Cell [11], so we are grateful to Sayan Bhattacharjee, Suvrajit Maji, Prikshat Dadhwal, Zhening Zhang, and Zuben P Brown for their work.
Competing interests
Columbia University has filed a patent application related to this work for which X.F. and J.F. are inventors.
Ethical considerations
No animal and/or human subjects were used in this study.
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Zheng, L., Liu, N., Gao, X., Zhu, W., Liu, K., Wu, C., Yan, R., Zhang, J., Gao, X., Yao, Y., et al. (2023). Uniform thin ice on ultraflat graphene for high-resolution cryo-EM. Nat Methods. 20(1): 123–130.
Article Information
Publication history
Received: Jul 9, 2024
Accepted: Dec 10, 2024
Available online: Jan 8, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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5,194 | https://bio-protocol.org/en/bpdetail?id=5194&type=0 | # Bio-Protocol Content
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Simple Method for Efficient RNA Extraction From Arabidopsis Embryos
FM Fernanda Marchetti
Gabriela Pagnussat
EZ Eduardo Zabaleta
In Press, Available online: Jan 05, 2025
DOI: 10.21769/BioProtoc.5194 Views: 25
Reviewed by: Noelia ForesiPooja Saxena Anonymous reviewer(s)
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Abstract
Plant embryos are contained within seeds. Isolating them is crucial when endosperm and seed coat tissues interfere with the study of mutant genetic functions due to differing genotypes between maternal and embryonic tissues. RNA extraction from plant embryonic tissue presents particular challenges due to the high activity of RNases, the composition of the seed, and the risk of RNA degradation. The developmental stage of the embryo is a key aspect of successful isolation and RNA extraction due to the size and amount of tissue. Proper handling during RNA extraction is critical to maintain RNA integrity and prevent degradation. While commercial kits offer various methods for RNA extraction from embryos, homemade protocols provide valuable advantages, including cost-effectiveness and accessibility for labs with limited funding. Here, we present a simple and efficient protocol for extracting RNA from isolated Arabidopsis thaliana embryos at the torpedo/cotyledon stage using a homemade extraction buffer previously reported for styles of Nicotiana alata.
Key features
• This straightforward homemade protocol builds upon established methods for embryo isolation [1] and RNA extraction [2,3].
• It enables the extraction of high-quality RNA from Arabidopsis embryos at the torpedo stage.
• The protocol is designed to be both simple and cost-effective, making it an excellent option for labs with limited funding.
Keywords: Arabidopsis embryo Embryo isolation RNA extraction Simple and efficient RNA extraction Homemade protocol Embryo development
Graphical overview
Simple method for RNA extraction from isolated Arabidopsis embryos using a homemade extraction buffer
Background
This protocol describes a straightforward two-step method for extracting RNA from isolated Arabidopsis embryos. Isolating RNA from plant tissues such as embryos, which are rich in RNases, poses significant challenges due to the risk of RNA degradation during sample preparation. To overcome these difficulties, commercial solutions such as RNAlater are commonly used to stabilize RNA by inactivating RNases, allowing for subsequent extraction using reagents like Trizol. In this protocol, we present a simple and effective homemade extraction buffer designed specifically for seed collection and RNA extraction. This buffer facilitates the isolation of high-quality RNA from Arabidopsis embryos. The method described here was successfully employed to extract RNA from developing embryos at the early torpedo and early cotyledon stages, from both immature (white) and mature (green) seeds. This approach offers an accessible and efficient alternative to commercial reagents, ensuring robust RNA recovery and preservation for downstream applications.
Materials and reagents
Biological materials
1. Arabidopsis thaliana ecotype Columbia (Col-0) wild type (WT) and embryo-lethal mutant plants from the Arabidopsis biological resource center (ARBRC, https://abrc.osu.edu/)
Reagents
1. Tris [Tri(hydroxymethyl)aminomethane] (Genbiotech, catalog number: RU2510)
2. EDTA (ethylenediamine tetraacetic acid disodium salt) (Biopack, catalog number: 1604.07)
3. Urea (Sigma, catalog number: U5378)
4. Sodium dodecyl sulfate (SDS) (Anedra, catalog number: 7211)
5. Ammonium acetate (Fluka, catalog number: 09690)
6. Lithium chloride (LiCl) (Sigma, catalog number: L9650)
7. 2-Mercaptoethanol pure (Biopack, catalog number: 9545.05)
8. Diethyl pyrocarbonate (DEPC) (Sigma, catalog number: D5758)
9. Percoll (GE Healthcare, catalog number: 17-0891-01)
10. DEPC (diethylpyrocarbonate) (Sigma, catalog number: D5758)
11. SYBRTM Safe DNA gel stain (Invitrogen, catalog number: S33102)
12. Agarose (Gene Biotech LE-Agarose 1200, catalog number: RU1010)
13. DEPC water (1 mL DEPC/L water; stirred overnight and autoclaved)
14. Phenol:chloroform:isoamyl alcohol 25:25:1 (Sigma, catalog number: P3803)
15. Chloroform (Biopack, catalog number: 1651.08)
16. Isopropanol (Mallinckrodt, catalog number: 3032-08)
Solutions
1. Buffer Tris HCl pH 8, 1 M (see Recipes)
2. EDTA pH 8, 0.5 M (see Recipes)
3. SDS 10% (see Recipes)
4. Ammonium acetate, 10 M (see Recipes)
5. Lithium chloride, 8 M (see Recipes)
6. Extraction buffer (see Recipes)
Recipes
1. Buffer Tris-HCl pH 8, 1 M
Weigh 12.11 g of Tris, add 75 mL of pure sterile water, adjust to pH 8 with HCl, and fill with pure sterile water to a final volume of 100 mL.
2. EDTA pH 8, 0.5 M
Weigh 18.6 g of EDTA, add 75 mL of pure sterile water, adjust to pH 8 using 10 M NaOH, and stir until EDTA is completely dissolved. Solubility improves while alkalinity increases; complete with pure sterile water to a 100 mL final volume.
3. SDS 10%
Dissolve 10 g of SDS in 100 mL of pure sterile water.
4. Ammonium acetate, 10 M
Dissolve 7.78 g of ammonium acetate in 10 mL of DEPC water.
5. Lithium chloride, 8 M
Dissolve 3.39 g of lithium chloride in 10 mL of DEPC water.
6. Extraction buffer (10 mL)
Reagent Final concentration Quantity or Volume
Urea 7 M 4.2 g
EDTA 10 mM 200 μL
Tris-HCl (1 M, pH 8) 100 mM 1 mL
SDS 10% 1% 1 mL
2-Mercaptoethanol 1% 100 μL
H2O to 10 mL *see note
*Note: Weigh urea in a 50 mL Falcon tube and dissolve urea in half volume (5 mL for a final volume of 10 mL) of water. Vortex the solution to help it dissolve. The bottom of smaller tubes hinders the full solubility of urea. Then, add the rest of the components and complete to final volume. Keep at room temperature. Cold will precipitate SDS in the extraction buffer. No risk for later RNA extraction.
Laboratory supplies
1. Syringe, needles (13 × 0.3 mm) and pointy tips tweezers (Dumont, catalog number: 0103-2-PO)
2. Plastic grinding rod for Eppendorf tubes (SSIbio, catalog number: 1005-39)
Equipment
1. Magnifying glass (LABKLASS, model: HG468405)
2. Table centrifuge (Eppendorf, model: centrifuge 5418)
3. Vortex (VornadoTM, Benchmark Scientific, catalog number/model: BV101-B)
4. Agarose gel electrophoresis equipment (Bio-Rad, model: PowerPacTM Basic Power Supply, Horizontal Electrophoresis Systems)
5. NanoDropTM One/One (Thermo Fisher, catalog number/model: ND-ONE-W)
Procedure
A. Embryo isolation
1. Collection of seeds
a. Add 100 μL of extraction buffer (see Recipes) in a 1.5 mL Eppendorf tube. Weigh the tube.
b. Using a needle under a magnifying glass, open mature or immature siliques to collect green and/or white seeds, respectively. Place the seeds inside the Eppendorf tube containing the extraction buffer. Collect seeds from approximately 25 siliques. Weigh the tube after collecting seeds. For a satisfactory RNA extraction, the minimum recommended amount for seed collection is 0.010 g of tissue.
c. Spin down the embryos in a table centrifuge at 1,700× g for 30 s. Carefully remove the extraction buffer by pipetting. Wash the embryos three times with 1 mL of DEPC water. Spin down at 1,700× g after each washing.
2. Embryo isolation from seed coat (from Perry and Wang [2])
a. Remove 750 μL of the DEPC water per tube.
b. Shake gently the lower part of the Eppendorf tube by hand (using fingers) just to let the seeds spread in the remaining water.
c. Use a plastic grinding rod for Eppendorf tubes (plastic sticks) to press softly the sample against the tube’s wall to release embryos from seeds by applying soft pressure. Repeat three times with smooth movements.
d. Transfer the sample (250 μL) by pipetting to a new tube containing 500 μL of DEPC water and 250 μL of Percoll (25% v/v Percoll). To facilitate pipetting the sample, use a 200 μL yellow tip with the tip cut off.
e. Centrifuge at 72× g for 10 min.
f. Remove the seed coats of the upper layer together with the Percoll solution by pipetting.
g. Resuspend the embryos carefully in 250 μL of the remaining Percoll solution and transfer the sample by pipetting to a new tube containing 0.75 mL of 25% v/v Percoll solution.
h. Centrifuge at 72× g for 10 min.
i. Remove the seed coats from the upper layer and discard the remaining Percoll by pipetting.
j. Wash the embryos three times with 1 mL of DEPC water. Spin down at 72× g after each washing.
B. RNA extraction [2,3]
Before starting, prepare Eppendorf tubes containing:
I. 500 μL of phenol:chloroform:isoamyl alcohol (25:24:1) + 500 μL of extraction buffer
II. 0.5 mL of phenol:chloroform:isoamyl alcohol (25:24:1)
III. 0.5 mL of chloroform
IV. 0.1 mL of 10 M ammonium acetate
1. Remove washing water from the tube by pipetting. Add 100 μL of extraction buffer and use a plastic grinding rod to crash the tissue against the tube wall. Crash embryos completely.
2. Add the sample to tube I and vortex immediately for 2 min.
3. Centrifuge at 18,000× g for 10 min at room temperature. Take the upper phase and transfer it to tube II. Vortex vigorously for 2 min.
4. Centrifuge at 18,000× g for 10 min at room temperature. Transfer the upper phase to tube III. Vortex vigorously for 2 min.
5. Centrifuge at 18,000× g for 10 min at room temperature. Transfer the aqueous phase to tube IV. Add 1 volume of cold isopropanol. Mix by inversion. Store at -20 °C for any duration between 30 min and overnight.
6. Centrifuge at 18,000× g for 15 min at 4 °C. Discard supernatant by inverting the tube. Dry pellet at room temperature.
7. Resuspend the pellet in 300 μL of DEPC water. Maintain tubes on ice. Check that the pellet is well-solubilized. Heat sample at 55 °C for 5 min to better solubilize the pellet (optional). Add 300 μL of 8 M LiCl and mix by inversion. Place the tubes inside an ice bucket and leave on ice overnight inside the refrigerator to precipitate RNA.
8. Centrifuge at 18,000× g for 15 min at 4 °C. Discard supernatant after centrifugation.
9. Resuspend the pellet in 300 μL of DEPC water. Add 100 μL of ammonium acetate. Fill the tube with 96% cold ethanol. Precipitate at -20 °C for at least 30 min.
10. Centrifuge at 18,000× g for 15 min at 4 °C. Wash pellet with 70% ethanol. Centrifuge at 18,000× g for 15 min at 4 °C. Discard supernatant. Dry the pellet at room temperature and resuspend in 20 μL of DEPC water.
11. Check RNA profile by loading 3 μL of RNA (approximately 100–150 ng of RNA) in a 1.5% agarose gel; run at 150 V for 30–40 min. RNA concentration yield is approximately 50–100 ng/μL RNA per 0.010 g of isolated seeds containing embryos at the torpedo stage.
12. RNA can be stored at -20 °C for short-term uses. Storage at -80 °C is recommended for long-term use.
Validation of protocol
This protocol was developed by combining two existing protocols:
For RNA extraction:
McClure et al. [2]. Self incompatibility in Nicotiana alata involves degradation of pollen rRNA. Nature.
Roldán et al. [3]. Molecular and genetic characterization of novel S-RNases from a natural population of Nicotiana alata. Plant Cell Rep (using extraction buffer, no commercial kits).
For embryo isolation:
Perry, S. E. and Wang, H. [1]. Rapid isolation of Arabidopsis thaliana developing embryos. BioTechniques. (using commercial stabilization solutions).
The innovation lies in the simultaneous application of both pre-existing protocols along with the incorporation of a homemade extraction buffer for RNA extraction from tissues with high RNase activity, such as Arabidopsis embryos, without relying on commercial kits. In our hands, the homemade extraction buffer effectively prevents RNA degradation from Arabidopsis embryos. The protocol presented here represents a valuable alternative for two challenging steps when working with Arabidopsis seeds: first, the separation of embryos from seed coats (embryo isolation step), and second, RNA extraction from tissues that require RNase inhibitors and stabilization solutions to prevent RNA degradation (RNA extraction using homemade buffer step).
General notes and troubleshooting
General notes
This protocol is an alternative for RNA extraction from isolated Arabidopsis embryos using a homemade extraction buffer to prevent RNA degradation of tissues with high RNAse activity.
Embryo developmental stages play a crucial role in obtaining the necessary amount of tissue. For example, mature seeds (light green) containing embryos at advanced developmental stages, such as early cotyledon or cotyledon stages, have larger embryos and provide higher amounts of tissue when collecting from Arabidopsis siliques. In contrast, immature seeds (white seeds) contain embryos at early torpedo and torpedo developmental stages, which are smaller than embryos found in mature seeds. However, 0.010 g of tissue yield around 100–150 ng/μL RNA, suitable for cDNA synthesis. Collection of 0.010 g can be performed in approximately 30 min. The more seeds collected, the more RNA can be obtained.
This protocol can be implemented not only for immature seeds containing embryos at the early torpedo stage but also for seeds at earlier developmental stages, for example, seeds that contain embryos at the heart stage. In those cases, collecting more tissue is required because embryos are smaller in size. In our hands, this protocol works great for the isolation of embryos at early stages of development, and RNA extraction succeeds. When collecting 0.010 g of immature seeds when working with embryos at the heart stage, the tissue yield is approximately 5–10 ng/μL RNA, also suitable for cDNA synthesis. However, when working with earlier embryo developmental stages, such as globular or early heart stages in which the embryonic amount of tissue is low (in relation to the whole seeds), this protocol can be applied by skipping the embryo isolation step to work with the whole seeds (embryo, endosperm, and seed coat), performing the step of RNA extraction straightforward with the homemade extraction buffer, which will prevent RNA degradation. When collecting 0.010 g of whole immature seeds, the yield is around 60–160 ng/μL RNA, which corresponds to RNA extracted from the entire seed and not only from isolated embryos as described above but also suitable for cDNA synthesis.
Measurements of RNA concentration were done using NanoDropTM by absorbance at 260 nm and purity was estimated by the ratio 260/280 nm. RNA profile was analyzed by running (approximately) 150 ng of RNA in a 1.5% agarose gel. Note that RNA extracted from plant “white” tissue, such as embryo or root, lacks plastid RNA. In these cases, the RNA profile is composed mainly of two bands corresponding to cytoplasmic 28S and 18S rRNA, different from the RNA profile extracted from leaves in which smaller bands are present, corresponding to organellar rRNAs.
In summary, the protocol presented here is a great alternative to extract RNA from isolated Arabidopsis embryos as well as directly from seeds (when containing embryos in early developmental stages). It may be very useful when studying, for example, embryo-lethal mutants. Besides, using a homemade buffer suitable to prevent RNA degradation represents a valuable alternative when there is limited access to commercial stabilization solutions or kits.
Acknowledgments
We thank CONICET and ANPCyT for funding as well as the Institute IIB-CONICET-UNMDP. We are grateful to Dr. Roldan for suggestions when performing the RNA extraction protocol.
Competing interests
There are no competing interests.
References
Perry, S. E. and Wang, H. (2003). Rapid isolation of Arabidopsis thaliana developing embryos. Biotechniques. 35(2): 278–282. https://doi.org/10.2144/03352bm06
McClure, B. A., Gray, J. E., Anderson, M. A. and Clarke, A. E. (1990). Self-incompatibility in Nicotiana alata involves degradation of pollen rRNA. Nature. 347(6295): 757–760. https://doi.org/10.1038/347757a0
Roldán, J. A., Quiroga, R. and Goldraij, A. (2010). Molecular and genetic characterization of novel S-RNases from a natural population of Nicotiana alata. Plant Cell Rep. 29(7): 735–746. https://doi.org/10.1007/s00299-010-0860-6
Article Information
Publication history
Received: Sep 17, 2024
Accepted: Dec 10, 2024
Available online: Jan 5, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
How to cite
Category
Plant Science > Plant molecular biology > RNA
Molecular Biology
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5,195 | https://bio-protocol.org/en/bpdetail?id=5195&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
A Bio-protocol resource
This is an In Press version of the protocol that has not yet been assigned to an issue.
Peer-reviewed
Colorimetric Determination of Tungsten and Molybdenum in Biological Samples
AD Aaron P. Donaghy
GS Gerrit J. Schut
NS Nana Shao
FP Farris L. Poole
Michael W. W. Adams
In Press, Available online: Jan 08, 2025
DOI: 10.21769/BioProtoc.5195 Views: 67
Reviewed by: Partha BasuKomuraiah MyakalaBhanu Jagilinki
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Abstract
Molybdenum (Mo) and tungsten (W) are elements that are utilized in biological systems. They are typically incorporated into the catalytic sites of enzymes coordinated to an organic pyranopterin cofactor; Mo may also be present in the form of a FeMo cofactor. While Mo is used by all branches of life, only a few microbes are able to utilize W. In order to study Mo- and W-dependent enzymes, it is important to be able to measure Mo and W in biological samples. Methods for determining Mo and W content in biological samples currently involve expensive and time-consuming processes like inductively coupled plasma mass spectrometry (ICP-MS) and chelation ion chromatography. There are less intensive colorimetric methods for measuring W in abiotic samples, but these have not been adapted to biological samples like cytosolic extracts and purified proteins. Herein, we developed a colorimetric assay based on the complexation of quercetin to molybdate (MoO42-) or tungstate (WO42-), the oxyanion forms of Mo and W that readily form in denatured biological samples. In the assay, the absorbance of quercetin is redshifted proportionally to the concentration of tungsten or molybdenum, which can be measured spectrophotometrically. This protocol provides a rapid method for screening biological samples for both Mo and W, although it does not distinguish between them.
Key features
• This protocol is adapted from the method developed by El-Sayed et al. [1] for analyzing abiotic samples.
• The protocol is designed for a 96-well plate format and optimized for analyzing protein samples.
• Can be used over the range of 1–20 μM W or Mo.
• Allows for rapid and high throughput Mo and W determination in samples during protein purification.
Keywords: Tungsten Molybdenum Colorimetry Quercetin UV-Vis Spectrophotometer Tungstoenzyme Molybdoenzyme Column Chromatography
Graphical overview
Overview of colorimetric determination of tungsten in protein samples using quercetin. Protein samples (200 μL) are transferred to microcentrifuge tubes along with 50 μL of 2.5× nitric acid mix and incubated at 65 °C overnight. The tubes are spun at 20,000× g for 20 min and 200 μL of the supernatant is transferred to a holding plate. 40 μL of each sample is transferred from the holding plate to four wells on an optical plate using a multichannel pipette. 160 μL of ethanol blank mix is added to one well per sample and 160 μL of quercetin mix is added to the remaining wells of each sample. The plate is sealed and shaken to mix each well. The seal is removed, and absorbances at 419 nm of each well are measured in a plate reader.
Background
Molybdenum (Mo) and tungsten (W) typically occur as pyranopterin cofactors at the catalytic sites of a variety of enzymes, except for Mo in the Mo-dependent nitrogenase, where it is part of a FeMo cofactor. Until now, methods for measuring these metals in biological samples were limited to resource-intensive protocols like high-performance liquid chromatography, ICP-MS, and voltammetry [2–4]. For the determination of Mo in the nitrogenase complex of Clostridium pasteurianum [5], a colorimetric assay was utilized, which involved the complexation and extraction of Mo with toluene-3,4-dithiol, determining Mo concentrations within the range of 0.2–100 μM [6]. However, this procedure is rather cumbersome and not readily adaptable to high throughput analysis. A simple colorimetric assay was developed for abiotic samples using a flavonol, quercetin, which complexes with free molybdate or tungstate in solution [1]. The formation of this complex is measured by its absorbance at 419 nm (Figure 1). In this report, the quercetin-based assay was adapted for biological samples, specifically protein samples from column chromatography fractions. Assay reagents and procedures were altered from the original assay to decrease interferences from substances present in biological samples, such as flavin and iron. To determine if the presence of protein, iron, and flavin in samples interfered with Mo and W measurements, standards were run with added protein (bovine serum albumin), iron (in the form of an iron-containing protein, a [4Fe-4S] ferredoxin), and flavin. As shown in Figures S1 and S2, the effects of these additions were small and not significant. The assay was also adapted to a 96-well plate format in order to simultaneously process large numbers of samples. Due to the chemical similarities between molybdate and tungstate, this assay can be utilized to determine the concentrations of either in biological samples but cannot distinguish between them [7].
Figure 1. Effect of tungsten and molybdenum on quercetin spectra. Graph illustrating the change in spectra (300–500 nm) of 1 mM quercetin without W or Mo (black), with 10 μM W (red), and with 10 μM Mo (blue) blanked against ethanol.
Materials and reagents
Biological materials
1. Native tungsten-containing proteins in the cytosolic extract of the anaerobic bacterium Eubacterium limosum were separated anaerobically using anion exchange chromatography and further purified using size exclusion chromatography (SEC) by a Bio-Rad Enrich SEC 650 (10 × 300) column using 25 mM Tris-HCl, pH 7.6, and 100 mM NaCl buffer. These fractions contain the tungsten-binding protein, Tub, which is a hexamer of 7 kDa subunits and binds a total of eight tungsten oxyanions [8]. Sixteen column fractions from the SEC column were used in the tungsten assay.
Reagents
1. Cetylpyridinium chloride (CPC) (Sigma-Aldrich, catalog number: C0732)
2. Deionized water (diH2O)
3. Ethanol, absolute (Decon Laboratories, catalog number: 2716)
4. 70% nitric acid (Fisher Chemical, catalog number: EW-88050-70)
5. Quercetin hydrate (Sigma-Aldrich, catalog number: 337951)
6. Sodium fluoride (Sigma-Aldrich, catalog number: S1504)
7. Sodium molybdate dihydrate (Sigma-Aldrich, catalog number: M1651)
8. Sodium tungstate dihydrate (Sigma-Aldrich, catalog number: S0765)
Solutions
1. 1 mM quercetin in ethanol
2. 10 mM CPC
3. 1 M nitric acid
4. 0.1% sodium fluoride
5. 100 μM sodium tungstate
6. 100 μM sodium molybdate
7. 2.5× nitric acid mix (see Recipes)
8. 1× nitric acid mix (see Recipes)
9. Ethanol blank mix (see Recipes)
10. Quercetin assay mix (see Recipes)
Recipes
1. 2.5× nitric acid mix
Reagent Final concentration Quantity or Volume
Nitric acid (1 M) 250 mM 25 mL
Sodium fluoride (0.1%) 0.0025% 2.5 mL
diH2O n/a 72.5 mL
Total n/a 100 mL
2. 1× nitric acid mix
Reagent Final concentration Quantity or Volume
Nitric acid (1 M) 100 mM 10 mL
Sodium fluoride (0.1%) 0.001% 1 mL
diH2O n/a 89 mL
Total n/a 100 mL
3. Ethanol blank mix
Reagent Final concentration Quantity or Volume
Ethanol (absolute) 10% 0.8 mL
Cetylpyridinium chloride (10 mM) 2.5 mM 2 mL
1× nitric acid mix n/a 5.2 mL
Total n/a 8 mL
4. Quercetin assay mix
*Note: Quercetin will begin to precipitate out of the solution more than 30 min after the formation of the mixture.
Reagent Final concentration Quantity or Volume
Quercetin (1 mM) in ethanol 0.1 mM 1.8 mL*
Cetylpyridinium chloride (10 mM) 2.5 mM 4.5 mL
1× nitric acid mix n/a 11.7 mL
Total n/a 18 mL
Laboratory supplies
1. 1.5 mL microcentrifuge tubes (Heathrow Scientific, catalog number: AXYRESV25S)
2. Plate-sealing film (EXCEL Scientific, Inc., catalog number: EZP-100)
3. 96-well assay microplates (Corning, catalog number: 9017)
4. Polypropylene 96-well plate (Elkay, catalog number: MICR-PVX)
5. PYREX® 50 mL round media storage bottles (Corning, catalog number: 1395-50)
6. PYREX® 100 mL round media storage bottles (Corning, catalog number: 1395-100)
7. Universal pipette tips, 200 μL (VWR, catalog number: 76322-516)
8. Universal pipette tips, 300 μL (VWR, catalog number: 76322-520)
9. Universal pipette tips, 1,250 μL (VWR, catalog number: 76322-138)
10. 50 mL polypropylene conical tubes (Geiner Bio-One, catalog number: 227 261)
Equipment
1. Centrifuge (Beckman Coulter Microfuge 20, catalog number: B31600)
2. Microplate reader (Molecular Devices, model: SpectraMax® 190)
3. Vortex mixer (Vortex-Genie 2, model: G-560)
4. P20 Pipetman (Gilson, catalog number: F123600)
5. P200 Pipetman (Gilson, catalog number: F123601)
6. P1000 Pipetman (Gilson, catalog number: F123602)
7. 5–100 μL 8-channel electronic pipette (Eppendorf, Xplorer®, catalog number: 4861000120)
8. 15–300 μL 8-channel electronic pipette (Eppendorf, Xplorer®, catalog number: 4861000147)
9. 20–1,200 μL 8-channel electronic pipette (Eppendorf, Xplorer®, catalog number: 4861000163)
Software and datasets
1. SoftMax® Pro 4.8, by Molecular Devices
2. Microsoft Excel
Procedure
A. Preparation of protein samples
1. Assemble up to 16 protein samples to test per plate.
2. Transfer 200 μL of each protein sample to labeled centrifuge tubes.
3. Add 50 μL of 2.5× nitric acid mix to each centrifuge tube.
4. Vortex tubes, seal, and incubate at 65 °C overnight.
5. Centrifuge tubes at 20,000× g for 20 min.
6. Transfer 200 μL of the supernatant from each tube to wells on a 96-well holding plate.
B. Preparation of standard
1. Prepare tungsten or molybdenum standard solutions by diluting 100 μM stock solution to produce concentrations of 0, 1, 2.5, 5, 7.5, 10, 15, and 25 μM.
2. Add 50 μL of 2.5× nitric acid mix to each standard solution.
3. Vortex solutions and transfer 200 μL of each solution to wells on a 96-well holding plate.
C. Plate assay
1. Transfer 40 μL of each sample and each standard to four wells on an optical 96-well plate.
2. Transfer 160 μL of ethanol blank mix to 1 out of 4 wells for each sample/standard.
3. Transfer 160 μL of quercetin mix to the three remaining wells for each sample/standard.
4. Place a plate seal on the plate and shake in a plate shaker to mix.
5. Remove the seal (see Troubleshooting 1) and measure absorbance at 419 nm in a plate reader.
Data analysis
The absorbance from the ethanol blank well of each sample is subtracted from the absorbances of the other three wells. The absorbance of the 0 μM tungstate in the standard is used as a quercetin blank and is subtracted from all absorbance values. The standard absorbance values are plotted against tungstate concentration to produce a standard curve (Figure 2). From the standard curve, an equation is generated by which the tungstate concentration in each sample well can be calculated and averaged.
Figure 2. Example standard curves for molybdenum (blue) and tungsten (red). Data points were corrected for background absorbance and zeroed against their respective 0 μM standards. Results show an average of three replicates. Error bars show calculated standard deviation. Limit of detection calculated from equation, (3.3/σregression)/slope, is 0.9 μM for W and 0.8 μM for Mo.
Validation of protocol
Native tungsten-containing proteins from Eubacterium limosum were separated using SEC, and the resulting fractions were tested using both the quercetin-linked assay protocol and ICP-MS. The latter technique was used to determine the accuracy of the colorimetric assay. For both procedures, three replicates were analyzed, and the results were averaged. The W concentrations for the quercetin assay matched well with the ICP-MS W values and resulted in a similar shape of the W peak from the SEC samples (Figure 3). Specifically, for fractions 16 and 17, the quercetin assay results correspond to those from ICP-MS analysis, with the former yielding concentrations that were 89% and 120%, respectively, of those determined by ICP-MS (Table 1). A tungsten concentration below ~0.25 μM in any fraction as determined by ICP-MS was not reliably detected with the quercetin assay. Nevertheless, the results from the quercetin assay correctly identified W-containing proteins in column chromatography fraction. The correlation between the two protocols was quantified with the production of a linear regression model using the results from each protocol. The model shows an R2 confidence value of 0.9373 (Figure S3).
Figure 3. Quercetin assay and ICP-MS results. Tungsten concentrations measured using the quercetin assay (red) and ICP-MS (black) [8] on SEC fractions containing the tungsten-storage protein, Tub. The ICP-MS protocol is described in Schut et al. [9]. For both the quercetin assay and ICP-MS, three replicates were averaged. Error bars show calculated standard deviation.
Table 1. Results for tungsten-containing fractions. Tungsten concentrations (μM) from fractions 15–20 calculated using the colorimetric quercetin assay and ICP-MS (control) along with percent recovery. For full sample set results, see Table S1.
Fraction # [Tungsten] (μM) – ICP-MS [Tungsten] (μM) – Quercetin assay % Recovery
15 0.17 0.01 3%
16 2.10 1.86 89%
17 0.94 1.13 120%
18 0.22 0.60 276%
19 0.06 0.00 0%
20 0.02 0.00 0%
General notes and troubleshooting
General notes
1. Due to the similarity in the chemical properties of molybdate and tungstate, this assay cannot distinguish between them. In samples with both molybdenum and tungsten, this assay will determine the sum of the concentrations of Mo and W in the sample.
2. Optimal W or Mo concentration range for the assay is from 1.0 μM to 20 μM. The detection limit calculated from the calibration curves (Figure 2) is ~0.8 μM for Mo and ~0.9 μM for W.
3. The presence of yellow-colored compounds that have absorbance near 420 nm in the samples may affect the calculated tungsten content. This issue can be mitigated by accounting for background interference using the ethanol blank mix.
Troubleshooting
Problem 1: Standard results produce poor linear regression line.
Possible cause: Bubbles in the wells have warped absorbance data.
Solution: After removing the plate seal, check to make sure no bubbles are present in the wells by gently tapping the plate against a flat surface.
Supplementary information
The following supporting information can be downloaded here:
1. Figure S1. The effect of protein, iron, and flavin on the molybdenum standard curve
2. Figure S2. The effect of protein, iron, and flavin on the tungsten standard curve
3. Figure S3. Correlation between quercetin assay and ICP-MS results
4. Table S1. Full 16-sample set results for tungsten determination
Acknowledgments
This work was funded by a grant from the National Institutes of Health (GM 136885).
This protocol is adapted from the method originally described by El-Sayed et al. [1].
Competing interests
The authors declare no competing interests.
Ethical considerations
No human or animal subjects were used in this protocol.
References
El-Sayed, A. A., Saad, E. A., Mohamed Ibrahime, B. M. and Tarek Mohamed Zaki, M. (2000). Flavonol Derivatives for Determination of Cr(III) and W(VI). Microchim Acta. 135: 19–27.
Bednar, A., Mirecki, J., Inouye, L., Winfield, L., Larson, S. and Ringelberg, D. (2007). The determination of tungsten, molybdenum, and phosphorus oxyanions by high performance liquid chromatography inductively coupled plasma mass spectrometery. Talanta. 72(5): 1828–1832.
Hagedoorn, P. L., van't Slot, P., van Leeuwen, H. P. and Hagen, W. R. (2001). Electroanalytical Determination of Tungsten and Molybdenum in Proteins. Anal Biochem. 297(1): 71–78.
Wang, J. (1992). Catalytic—adsorptive stripping voltammetric measurements of ultratrace levels of tungsten. Talanta. 39(7): 801–804.
Vandecasteele, J. P. and Burris, R. H. (1970). Purification and Properties of the Constituents of the Nitrogenase Complex from Clostridium pasteurianum. J Bacteriol. 101(3): 794–801.
Clark, L. J. and Axley, J. H. (1955). Molybdenum Determination of Soils and Rocks with Dithiol. Anal Chem. 27(12): 2000–2003.
Mohamed Zaki, M. T. and El-Sayed, A. A. (1992). Use of quercetin in an improved method for molybdenum determination in waste water, silicate rocks and diverse alloys. Mikrochim Acta. 106: 153–161.
Shao, N., Dayong, Z., Schut, G. J., Poole, F. L., Coffey, S. B., Donaghy, A. P., Putumbaka, S., Thorgersen, M. P., Chen, L., Rose, J., et al. (2024). Storage of the Vital Metal Tungsten in a Dominant SCFA-Producing Human Gut Microbe Eubacterium limosum and Implications for Other Gut Microbes.
Schut, G. J., Thorgersen, M. P., Poole, F. L., Haja, D. K., Putumbaka, S. and Adams, M. W. W. (2021). Tungsten enzymes play a role in detoxifying food and antimicrobial aldehydes in the human gut microbiome. Proc Natl Acad Sci USA. 118(43): e2109008118.
Article Information
Publication history
Received: Oct 28, 2024
Accepted: Dec 16, 2024
Available online: Jan 8, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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5,196 | https://bio-protocol.org/en/bpdetail?id=5196&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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Peer-reviewed
Automated Sequential Derivatization for Gas Chromatography-[Orbitrap] Mass Spectrometry-based Metabolite Profiling of Human Blood-based Samples
AJ Akrem Jbebli
KC Kateřina Coufalíková
MZ Moira Zanaboni
MB Manuela Bergna
RP Renzo Picenoni
JK Jana Klánová
EP Elliott J. Price
In Press, Available online: Jan 08, 2025
DOI: 10.21769/BioProtoc.5196 Views: 153
Reviewed by: Amit Kumar DeyJoyce Chiu Anonymous reviewer(s)
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Abstract
Many small molecules require derivatization to increase their volatility and to be amenable to gas chromatographic (GC) separation. Derivatization is usually time-consuming, and typical batch-wise procedures increase sample variability. Sequential automation of derivatization via robotic liquid handling enables the overlapping of sample preparation and analysis, maximizing time efficiency and minimizing variability. Herein, a protocol for the fully automated, two-stage derivatization of human blood–based samples in line with GC–[Orbitrap] mass spectrometry (MS)-based metabolomics is described. The protocol delivers a sample-to-sample runtime of 31 min, being suitable for better throughput routine metabolomic analysis.
Key features
• Direct and rapid methoximation on vial followed by silylation of metabolites in various blood matrices.
• Measure ~40 samples per 24 h, identifying > 70 metabolites.
• Quantitative reproducibility of routinely measured metabolites with coefficients of variation (CVs) < 30%.
• Requires a Thermo ScientificTM TriPlusTM RSH (or comparable) autosampler equipped with incubator/agitator, cooled drawer, and automatic tool change (ATC) station equipped with liquid handling tools.
Keywords: Automation Derivatization Thermo ScientificTM TriPlusTM RSH Metabolite profiling Gas chromatography–mass spectrometry (GC–MS)
Graphical overview
Workflow for profiling metabolites in human blood using automated derivatization
Background
Gas chromatography–mass spectrometry (GS–MS)-based blood metabolite profiling is an established biochemical technique to investigate human disease and treatment response [1,2], being scalable to large epidemiological studies [3]. Most analysis is conducted on plasma and serum routinely collected in clinical settings, but the use of dried blood spots is prevalent for newborn screening [4] and forensic toxicology, increasingly being collected outside clinical settings for population studies [5].
Once samples are in the lab, sample preparation often constitutes more than 60% of total analytical time [6,7], representing the major bottleneck of throughput. Manual handling during sample preparation contributes to higher bias, with increasing sample-to-sample variation that affects the results and biological interpretation [8].
Derivatization is often employed to make polar non-volatile compounds amenable to GC–MS analysis [9,10]. In human studies, the most used derivatization method for the detection of metabolites is methoximation followed by silylation. Methoximation stabilizes the carbonyl group of, e.g., reducing sugars, whilst silylation acts upon a diversity of functional groups (alcohols, carboxylic acids, amines, thiols, and phosphates), enabling the detection of a broad range of metabolites. Additionally, available electron ionization (EI) MS libraries comprise a high proportion of spectra for trimethylsilyl (TMS) derivatives [11,12] to aid annotation.
However, classical batch-wise derivatization approaches often lead to high sample variability, particularly because of the differing length of time following derivatization until injection. Notably, many derivatization reactions do not have a defined endpoint, and derivatives are often unstable. Furthermore, many analytes form multiple derivatives and isomers [13,14], complicating comparative quantification due to inconsistent ratios of different derivative forms.
To overcome the variability of batch-wise derivatization, procedures for automated, sequential derivatization for human plasma GC–MS metabolomics have been described [15,16]. However, both reported applications had relatively long GC runtimes (>40 min) and sample-to-sample runtimes of ~1 h or more. Additionally, their application was not demonstrated for other blood matrices.
Herein, we describe an automated method for the methoximation-silylation derivatization of metabolites for the sequential, online GC-[Orbitrap] MS-based metabolite profiling of various dried blood matrices, i.e., dried blood spots of capillary and venous blood, dried serum and plasma, and synthetic blood. The sequential sample preparation standardizes the time from derivatization to sample injection, increasing robustness over batch-wise procedures [17], and provides reproducible measurements of ~70 confirmed metabolites per sample with a throughput of ~45 injections per 24 h.
Materials and reagents
Biological materials
1. Dried capillary blood spots (DBS capillary): Capillary blood was collected using a lancet from the finger and spotted onto Whatman 903 protein saver cards [18]. DBS-capillary spots were dried for 12 h at 20 °C in a fume hood. Dried spots were covered with aluminum foil, placed in a zip-lock bag with silica bags to absorb moisture, and stored at -80 °C
2. Dried venous blood spots (DBS venous): Venous blood was collected via S-Monovette® neutral Z (Sarstedt, catalog number: 04.1926.001) [18]. DBS-venous spots were made by dispensing 100 μL onto collection cards following the same procedure as for DBS capillary
3. Gibco human plasma-like material (HPLM) (Thermo Scientific, catalog number: A4899101)
4. Pooled human serum: Mixture of serum derived from patients of University Hospital Brno
5. Pooled human plasma: Mixture of plasma derived from patients of University Hospital Brno
Reagents
1. Chloroform (Sigma-Aldrich, catalog number: 65049)
2. Isooctane (Sigma-Aldrich, catalog number: 32291-M)
3. Methanol (Biosolve, catalog number: 136841)
4. Hexane (J.T. Baker, catalog number: 5274.2500)
5. Methoxylamine hydrochloride (Thermo Scientific, catalog number: A19188)
6. N-Methyl-N-trimethylsilyltrifluoroacetamide (MSTFA), ≥98.5% (Sigma-Aldrich, catalog number: 69479-25ML)
7. Pyridine anhydrous, 99.8% (Sigma-Aldrich, catalog number: 270970-100ML)
8. Acetone, ≥99.5% (J.T. Baker, catalog number: 15548454)
9. C7-C40 saturated alkanes standard, certified reference material (CRM) (MilliporeSigmaTM SupelcoTM, catalog number: 11-101-7207)
10. Fully resolved native Mono-Deca PCB mixture (unlabeled) (Cambridge Isotope Laboratories, Inc., catalog number: EC-5434)
11. D7-cholesterol, >99% (Sigma-Aldrich, catalog number: 700041P-10MG)
12. D5-glycine, 99% (Sigma-Aldrich, catalog number: 175838-5G)
13. D27-myristic acid, 99% (Sigma-Aldrich, catalog number: 366889-1G)
Solutions
1. C7-C40 alkanes solution (1 μg/mL) (see Recipes)
2. Mono-Deca PCBs solution (100–200 ng/mL) (see Recipes)
3. D7-Cholesterol stock solution (100 μg/mL) (see Recipes)
4. D5-Glycine stock solution (100 μg/mL) (see Recipes)
5. D27-Myristic acid stock solution (100 μg/mL) (see Recipes)
6. Internal standard mix (IS mix, 1.6 μg) (see Recipes)
7. Methoximation solution (MeOx, 15 mg/mL, 1 μg/mL ISs) (see Recipes)
8. MSTFA working solution (see Recipes)
Recipes
1. C7-C40 alkanes solution (1 μg/mL)
The CRM C7-C40 saturated alkanes standard contains each alkane at 1,000 μg/mL in hexane. The mixture is diluted in isooctane to achieve a final concentration of 1 μg/mL per each alkane component in a 1 mL final solvent volume. Stored at -20 °C, the solution should be stable for up to a year. However, due to multiple freeze-thaw cycles, we recommend preparing fresh solution every 6 months.
2. Mono-Deca PCBs solution (100–200 ng/mL)
The fully resolved native Mono-Deca PCB mixture (unlabeled) is composed of components at 1,000–2,000 ng/mL in isooctane. The mixture is diluted 10-fold to give a solution of components at 100–200 ng/mL in 1 mL of isooctane. Stored at -20 °C, the solution should be stable for up to a year. However, due to multiple freeze-thaw cycles, we recommend preparing fresh solution every 6 months.
3. D7-Cholesterol stock solution (100 μg/mL)
D7-Cholesterol stock solution (100 μg/mL) was prepared by dissolving 10 mg of D7-Cholesterol in 1 mL of chloroform, aliquoting 100 μL into 2 mL amber vials (i.e., 100 μg aliquots), and drying for 20 min via centrifugal evaporator operating the low boiling point (bp) program for solvents bp 60–100 °C, without light. Prior to use, 100 μg/mL stock solution is made by reconstitution in 1 mL of chloroform.
4. D5-Glycine stock solution (100 μg/mL)
D5-Glycine stock solution (100 μg/mL) is prepared by dissolving 10 mg of D5-Glycine in 1 mL of methanol, aliquoting 100 μL into 2 mL amber vials (i.e., 100 μg aliquots), and drying for 20 min via centrifugal evaporator operating the low boiling point (bp) program for solvents bp 60–100 °C, without light. Prior to use, 100 μg/mL stock solution is made by reconstitution in 1 mL of methanol.
5. D27-Myristic acid stock solution (100 μg/mL)
D27-Myristic acid stock solution (100 μg/mL) is prepared by dissolving 10 mg of D27-Myristic acid in 1 mL of chloroform, aliquoting 100 μL into 2 mL amber vials (i.e., 100 μg aliquots), and drying for 20 min via centrifugal evaporator operating the low boiling point (bp) program for solvents bp 60–100 °C, without light. Prior to use, 100 μg/mL stock solution is made by reconstitution in 1 mL of chloroform.
6. Internal standard mix (IS mix, 1.6 μg)
Internal standard mix is prepared by dispensing 16 μL of each prepared 100 μg/mL stock standard solution (D7-Cholesterol, D5-Glycine, and D27-Myritic acid) in 2 mL amber vials. The mix is evaporated to dryness via centrifugal evaporator using the same conditions as previously mentioned (~20 min, no light, and low bp program). Vials are capped with the bonded PTFE/silicone septum screw cap and stored at -80 °C.
Note: Each individual labeled standard stock solution and the IS mix listed above are stored dried at -80 °C; in our experience, they remain stable for >6 months. We recommend periodic monitoring by GC analysis to check stability.
7. MeOx solution (15 μg/mL, 1 μg/mL ISs)
A MeOx stock (15 μg/mL) is freshly prepared every two days via weighing 150 mg of methoxylamine hydrochloride into a 10 mL amber flat-bottom glass vial and dissolving in 10 mL of pyridine. For use, 1.6 mL of MeOx stock is dispensed into the IS mix vials (2–3), capped with pre-slit caps, and vortexed (10 s, 2,000 rpm) to give MeOx solution (15 μg/mL, 1 μg/mL ISs). MeOx solution (15 μg/mL, 1 μg/mL ISs) is placed in the cooled drawer of the autosampler with a controlled temperature at 5 °C and sealed under nitrogen.
8. MSTFA working solution
1.6 mL of MSTFA is dispensed in 2 mL amber vials (2–3) capped with bonded pre-slit PTFE/silicone septum screw caps and placed in the cooled drawer.
Laboratory supplies
1. 2 mL amber vial (Agilent, catalog number: 5190-9063)
2. Bonded PTFE/silicone septum screw cap (Agilent, catalog number: 5190-9068)
3. Magnetic cap Silicone/PTFE/starburst-slitted septa (PAL, catalog number: Cap-ND9-St-SP10Sb-100)
4. Glass amber vial with 0.2 mL integrated insert (MACHEREY-NAGEL, catalog number: 702008)
5. 10 mL amber flat bottom glass vial (JG Finneran, catalog number: 31018F-2346A) with Screw cap, solid top with PTFE liner (Supelco, catalog number: 27163)
6. Bonded pre-slit PTFE/silicone septum screw cap (Agilent, 5185-5824)
7. 0.1–10 μL epT.I.P.S. Reloads (Eppendorf, catalog number: 022491504)
8. 2–200 μL epT.I.P.S. Reloads (Eppendorf, catalog number: 022491733)
9. 50 μL–1 mL epT.I.P.S. Reloads (Eppendorf, catalog number: 022491555)
10. 0.5–10 mL epT.I.P.S. Standard (Eppendorf, catalog number: 022492098)
11. Labels for vials
12. Whatman 903 cards (Merck, catalog number: WHA10531018)
Equipment
1. 0.5–10 μL pipette (Eppendorf Research® plus, catalog number: 3123000020)
2. 20–200 μL pipette (Eppendorf Research® plus, catalog number: 3123000055)
3. 100–1,000 μL pipette (Eppendorf Research® plus, catalog number: 3123000063)
4. 1,000–10,000 μL pipette (Eppendorf Research® plus, catalog number: 3123000080)
5. 3 mm hole puncher
6. Centrifugal evaporator (SP Genevac EZ-2 Series centrifugal evaporator, model: 3.0, Elite)
7. Storage boxes for 2 mL vials (VWR, catalog number: 525-0934)
8. Mini vortex (Wizard D, catalog number: 444-0746)
9. AS X2 PLUS analytical balance (RADWAG, product code: WL-104-0191)
10. O-rings, Viton (Restek, catalog number: 22242)
11. Thermo ScientificTM TriPlusTM RSH with extended rail (Thermo, catalog number: 1R77010-0400)
12. Thermo Scientific OrbitrapTM ExplorisTM GC 240 mass spectrometer (Thermo, catalog number: BRE725537)
13. TriPlus RSH, automatic tool change (ATC) station (Thermo, catalog number: 1R77010-1019)
14. 2 TriPlus RSH, liquid tool: D7, 57 mm (Thermo, catalog number: 1R77010-1007)
15. Triplus RSH autosampler syringe 57 mm, 23 s Ga, cone, 100 µL (Thermo, catalog number: 365H2141)
16. TriPlus RSH autosampler syringe 57 mm, 23s Ga, cone, 10 µL (Thermo, catalog number: 365D0311)
17. TriPlus RSH, agitator/incubator (Thermo, catalog number: 1R77010-1032)
18. TriPlus RSH, temperature-controlled drawer: single (Thermo, catalog number: 1R77010-1028)
19. Topaz Liner, splitless, gooseneck. w/wool 4 mm × 6.3 × 78.5 (Restek, catalog number: 23303)
20. Merlin microseal (Restek, catalog number: 22812)
21. Rxi-5Sil MS, 0.25 μm, 0.25 mm ID low polarity phase, 15 m length, 0.25 mm ID, 0.25 μm film thickness (Restek, catalog number: 13620)
22. Rxi guard column phase free, 0.53 mm ID, 2 m length to inlet (Restek, catalog number: 10073)
23. Rxi guard column phase free, 0.25 mm ID, 2 m length to spectrometer (Restek, catalog number: 10059)
24. Tweezers
Software and datasets
1. Sampling Workflow Editor (SWE) (version 1.4.0.4)
2. Thermo Scientific Xcalibur (version 4.4.16.14, February 6, 2020)
3. Chromeleon 7.2.10
4. PALscript Editor (version 3.1 Beta)
5. EIRENE-CZ_RECETOX Metabolome high resolution–electron ionization–mass spectral library (HR-[EI+]-MS), free, CC-BY-NC, https://doi.org/10.5281/zenodo.5483565
6. MS-DIAL (version 4.9.221218), free, CC-BY 4.0, https://doi.org/10.5281/zenodo.12589462
7. Skyline (version 23.1.0.268), free, modified Apache 2.0 License, https://github.com/ProteoWizard/pwiz/tree/master/pwiz_tools/Skyline
8. Spreadsheet software, e.g., Microsoft Excel, Google Sheets (free under Google terms of service)
9. BioRender (https://www.biorender.com/). The following figures were created using BioRender: Graphical overview, https://BioRender.com/v67e825; Figure S1, https://BioRender.com/y04y703
Procedure
A. Sample preparation: manual steps
Sample preparation of dried blood spots
1. Punch 3.3 mm from the DBS cards.
2. Place the punch using tweezers directly in a glass amber vial with 0.2 mL integrated insert and seal with magnetic cap silicone/PTFE/starburst-slitted septa.
Sample preparation of liquid blood matrices
3. Homogenize samples by gentle shaking and pipetting with ~5 inversions and 3 times pipetting (serum/plasma) or vortexing for ~10 s (HPLM).
4. Dispense 3 μL of liquid blood matrices in a glass amber vial with 0.2 mL integrated insert and seal with magnetic cap silicone/PTFE/starburst-slitted septa.
5. Dry for 20 min using the centrifugal evaporator, with the aqueous program with the lamp off.
Sample preparation of blanks
6. Prepare procedural blanks for DBS analysis by punching 3.3 mm from unused DBS cards into glass amber vials with 0.2 mL integrated insert and seal with magnetic cap silicone/PTFE/starburst-slitted septa. Use empty vials as procedural blanks for liquid blood samples and cap with magnetic cap silicone/PTFE/starburst-slitted septa.
7. Place vials containing samples, procedural blanks, and reagents (pyridine, Mono-Deca PCBs solution, C7-C40 alkane solution, MeOx solution, and MSTFA working solution) in the cooled drawer tray. Figure S1 shows an example of the tray layout.
Note: Pyridine is used as instrumental blank; the Mono-Deca PCBs solution is used for system suitability testing, e.g., checking mass accuracy, retention, and ion abundance prior to analysis; and C7-C40 alkane solution is used to generate external non-isothermal Kováts retention index [19]. These solutions are directly injected, i.e., without automated sample preparation, as shown in Table S1.
B. Automated sample preparation
The automated sample preparation is performed on a Thermo ScientificTM TriPlusTM RSH equipped with modules as displayed in Figure S2.
1. The automated method was created using SWE as detailed in standard operating procedure (SOP) (https://doi.org/10.5281/zenodo.10612856) and edited according to SOP (https://doi.org/10.5281/zenodo.10612909). The finalized method script is available in File S1, and a video simulating the method is available in File S2. Upload the method to the analytical sequence.
2. Run the sequence containing details about the injection list, instrumental method, sample position, and injection volume. Table S1 shows the sequence layout.
C. GC-HRMS analysis
Once sample preparation finishes, 1 μL of the derivatization product is automatically injected into the chromatographic system in splitless mode. The gas chromatography and mass spectrometry acquisition parameters are the same as previously provided [20,21].
In brief, analytes are separated upon a Restek Rxi5-Sil MS column (15 m, 0.25 mm ID, 0.25 μm) connected to a 2 m pre- and post-column. The GC oven temperature is initially set to 80 °C for 0.5 min, ramped to 200 °C at 40 °C/min with a hold time of 0.5 min¸ followed by a second ramp of 40 °C/min to 260 °C (also with 0.5 min hold), and then a third ramp of 55 °C/min to 330 °C with 4 min hold. The auxiliary temperatures are 280 °C. Helium is used as carrier gas at 1.2 mL/min. Electron ionization is performed at 70 eV with 50 μA emission current and 15 V electron lens. Data are acquired in profile mode, scanning 70–700 m/z at a resolving power of 60 K at 200 m/z. The SSL injector is equipped with a Merlin micro-seal and topaz liner. Exchange the topaz liner and O-ring for each sequence.
Data analysis
A. Metabolite composition analysis
1. Non-targeted data processing is performed in MS-DIAL [22,23]. Settings for detection (peak picking), deconvolution, alignment, and identification for MS-DIAL are provided in Table S2. Import data in .raw profile format.
2. Manually check peak integrations in MS-DIAL and update where needed.
3. Export alignment results and select raw data matrix (area) and representative spectra.
4. Open the peak area report as a spreadsheet (.csv) and, per each feature, subtract the maximum value detected in any procedural blank from the respective samples measured in the same analytical batch. Subsequent analysis can be performed on the finalized peak area matrix and/or mass spectra (.msp).
B. Selected metabolite quantification
1. Peak picking and integration are performed using the molecule interface of Skyline [24,25]. Transition settings and molecule settings are provided in Table S3 and Table S4, respectively.
2. A transition list of 52 TMS derivatives, representing 34 selected metabolites plus the 3 deuterated internal standards, is provided in Table S5 for targeted integration. The transition list contains at least three ions selected per analyte. Ions and retention time windows were selected based upon analysis of reference standards per each analyte [included in the EIRENE-CZ_RECETOX metabolome high resolution–electron ionization–mass spectral library (HR-[EI+]-MS)]. When possible, the theoretical masses of ions are used based on fragment formula annotation [26].
3. Import data in .raw format, manually check, and update peak integrations.
4. Export report for molecular transition results containing integrated peak areas per ion.
5. Open the molecular transition report as a spreadsheet (.csv) and select a quantitative ion per metabolite, e.g., select the ion showing the lowest coefficient of variation (CV).
6. Sum the integrated peak areas of the chosen quantification ions for all derivatives of an analyte [14].
7. Calculate the CVs per analyte quantitation based on the summed data. Subsequent analysis can be performed on the finalized metabolite area dataset.
Validation of protocol
The method was applied to the various blood samples, i.e., DBS capillary, DBS venous, serum, and plasma, with each matrix being analyzed in six replicates. For DBS samples, the six punches came from six different spots collected from an individual’s single timepoint blood draw. Raw files and Skyline processing are available on Panorama Public [27,28] at https://doi.org/10.6069/aeg8-9065.
The average CV across blood sample types was 2% for D27-myristic acid (1TMS derivative), 2% for D5-glycine (3 TMS derivative), and 4% for D7-cholesterol (1TMS derivative), evidencing high reproducibility of the automated sample preparation.
Analysis of generated data via MS-DIAL led to the detection of >600 deconvoluted features per matrix. Using a threshold of 30% CV [3], 449, 450, 264, and 329 features are reproducibly measured in serum, plasma, DBS venous, and DBS capillary, respectively (Figure 1). Over 70 analytes had confirmed identification for each matrix, i.e., through comparison with the RECETOX Metabolome HR-[EI+]-MS library (Table 1, Table S6). Analytes reported with higher confidence in annotation showed lower relative standard deviation across replicates, likely owing to a positive correlation between analyte abundance, peak, and spectral quality.
Table 1. Summary of deconvoluted features per blood matrix
Sample Features number All features mean average CV (%) Identified* analytes Identified analytes mean average CV (%)
Serum 839 39 77 22
Plasma 889 40 75 19
DBS venous 626 43 70 25
DBS capillary 650 43 69 23
*Analytes are annotated by comparison with RECETOX Metabolome HR-[EI+] MS library.
Figure 1. Violin plots of the coefficients of variation (CV) for peak area of features detected following GC-Orbitrap MS metabolomics analysis of various blood samples. Serum, plasma, DBS capillary, and DBS venous samples were analyzed in six replicates. A greater proportion of features are reproducibly measured, i.e., below the 30% CV threshold (dashed line), in liquid blood samples. The median is indicated by a white dot and the interquartile range (IQR) by a black box.
A selective target integration of a subset of 34 confirmed analytes also present in HPLM [as a surrogate certified reference material (CRM) material] was performed via Skyline. Multiple features derived from a single analyte are summed in order to check quantitative reproducibility [14]. The CVs for metabolites are below 30% except for asparagine, creatinine, taurine, and urea in serum; histidine in plasma; and creatinine, histidine, hypoxanthine, and taurine in HPLM (Figure 2).
Figure 2. Radar chart showing the coefficients of variation (CVs) of selected metabolites quantified by GC-Orbitrap MS metabolomics analysis of various blood samples. Serum, plasma, DBS capillary, DBS venous, and HPLM samples were analyzed in six replicates. When analytes have multiple derivatives, peak areas have been summed. Alanine (Ala), Arginine (Arg), Asparagine (Asn), Aspartic acid (Asp), Ornithine (Orn), Citric acid (Cit), Cysteine (Cys), Glutamic acid (Glu), Glycine (Gly), Histidine (His), Hydroxyproline (Hyp), Isoleucine (Ile), Glutamine (Gln), Leucine (Leu), Lysine (Lys), Threonine (Thr), Methionine (Met), Phenylalanine (Phe), Proline (Pro), Serine (Ser), Taurine (Tau), Tryptophan (Trp), Tyrosine (Tyr), Valine (Val), Creatinine (Crn), Fructose (Fru), Glucose (Glc), Galactose (Gal), Hypoxanthine (Hpx), Malic acid (Mal), Myoinositol (Myo), Niacinamide (Nam), Succinic acid (Suc), Urea (Urea).
General notes and troubleshooting
General notes
1. The workflow of sampling preparation was created and edited using Sampling Workflow Editor; however, the workflow was further optimized and improved using PALscript Editor.
2. Thermo ScientificTM GC-MS system equipped with Thermo ScientificTM TriPlusTM RSH needs to be configured for manipulation under Xcalibur or Chromeleon before running sample preparation and analysis (for that, use Thermo Foundation Instrument to configure the different compartments of the instrument under Xcalibur or Chromeleon Services Manager to configure under Chromeleon).
3. The derivatization method can also be applied to dried extracts and other materials, e.g., dried urine and seminal plasma samples (as per https://doi.org/10.5281/zenodo.7462217 [21] and https://doi.org/10.5281/zenodo.5734331 [20], respectively).
4. Alternatively, internal standards can be added directly to the vial containing MeOX solution. We used D5-glycine, D27-myristic acid, and D7-cholesterol because they elute at the beginning, middle, and end of the chromatogram and represent different compound classes.
5. The blood collected here was from adults, but collection can be applied to children and neonates. Different DBS sampling cards can be used.
6. HPLM was used as a surrogate certified reference material, but alternative materials can be used, e.g., NIST Standard Reference Material (SRM) 1950.
7. Tutorial for MS-DIAL data processing: https://systemsomicslab.github.io/mtbinfo.github.io/MS-DIAL/tutorial
8. Tutorial for Skyline (Hi-Res Metabolomics) data processing: https://skyline.ms/wiki/home/software/Skyline/page.view?name=tutorial_hi_res_metabolomics
9. Other libraries can be used for spectral matching, e.g., NIST/EPA/NIH Mass Spectral Library (proprietary).
10. The Thermo ScientificTM TriPlusTM RSH protocol can be coupled to the analysis of sample extracts via any coupled GC–MS model (e.g., single quadrupole, time of flight), yet analyte coverage and detection will depend on GC–MS methodology.
11. The procedure can be adapted to comparable cartesian autosamplers (e.g., PAL RTC, Gerstel MPS) if parameters can be mapped to the respective instrument control software.
Troubleshooting
1. We recommend the use of PAL magnetic caps to avoid collisions.
2. If 2 mL vials are transported, it is recommended to remove the magnetic transport ring (adapter) that is used in the case of 20/10 mL vials.
3. Cleaning the syringe needle and plunger used for sample preparation (here, 100 μL syringe) is recommended after running the sequence to avoid crystallization of the reagent. Wipe the plunger clean with a lint-free tissue, taking care not to bend the plunger. Remove the plunger and fill the syringe with solvent, insert the plunger back, and gently push the solvent through the needle. In this application, methanol and acetone were used as solvents for cleaning.
4. Pre-slitted caps are recommended for samples and reagents that will be penetrated multiple times.
Supplementary information
The following supporting information can be downloaded here:
1. Figure S1. Layout of reagents and samples in VT 54 cooled drawer trays. Example of sequence is provided in Table S1 for 24 h operation. Created in BioRender. Omics, M. (2024) https://BioRender.com/y04y703.
2. Figure S2. Thermo ScientificTM TriPlusTM RSH autosampler configuration. 1). X-axis extended rail with built in electronic and control. 2). Autosampler’s head with Z-axis. 3). Automatic tool change (ATC) station equipped with liquid handling tools. 4). Large wash station: 2 × 200 mL vials and a waste port. 5). Cooler drawer for controlled cool temperature and sealed under nitrogen: 2 × VT54 trays. 6). Standard wash station: 5 × 10 mL vials. 7). Agitator with controlled temperature for incubation with gentle shaking 6 positions adaptors installed for 2 mL vials.
3. Table S1. Example sequence layout for 24 h operation.
4. Table S2. MS-DIAL parameters.
5. Table S3. Skyline transition settings.
6. Table S4. Skyline molecule settings.
7. Table S5. Skyline transitions list to quantify select metabolites.
8. Table S6. List of analytes identified in blood samples.
9. File S1. Autosampler method script (.xml).
10. File S2. Video showing simulated sample preparation.
Acknowledgments
This work was supported by RECETOX research infrastructures (No. LM2023069) and financed by the Czech Ministry of Education, Youth and Sports. The work was financed from project SALVAGE (CZ.02.01.01/00/22_008/0004644) financed by MEYS – Co-funded by the European Union. This work was also supported by the European Union’s Horizon 2020 research and innovation program under grant agreement No 857560 (CETOCOEN Excellence). This publication reflects only the author's view, and the European Commission is not responsible for any use that may be made of the information it contains. The authors also wish to acknowledge Hagen Gegner for his thorough proofreading and for the feedback and insightful suggestions that greatly improved the quality of the manuscript. We thank Dalibor Valík for provision of pooled blood materials.
Competing interests
Moira Zanaboni and Manuela Bergna are employees of Thermo Fisher Scientific S.p.A. Renzo Picenoni is an employee of CTC Analytics AG.
Ethical considerations
The collection of pooled blood materials was acquired from studies involving humans approved by the University Hospital Brno, Multicentric Ethics Review Board, approval No. 03-091122/EK. The studies were conducted in accordance with the local legislation and institutional requirements. Samples used in this study were acquired from a by-product of routine care or industry. Written informed consent for participation was not required from the participants or the participants’ legal guardians/next of kin in accordance with the national legislation and institutional requirements.
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Article Information
Publication history
Received: Sep 12, 2024
Accepted: Dec 12, 2024
Available online: Jan 8, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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5,197 | https://bio-protocol.org/en/bpdetail?id=5197&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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Peer-reviewed
Profiling the Secretome of Glioblastoma Cells Under Histone Deacetylase Inhibition Using Mass Spectrometry
AM Aline Menezes
YM Yara Martins
FN Fábio César Sousa Nogueira
DP Denise de Abreu Pereira
KC Katia Carneiro
Published: Vol 15, Iss 3, Feb 5, 2025
DOI: 10.21769/BioProtoc.5197 Views: 110
Reviewed by: Nona FarbehiRupam Ghosh Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Molecular & Cellular Proteomics Mar 2024
Abstract
Glioblastoma (GBM) is the most aggressive brain tumor, and different efforts have been employed in the search for new drugs and therapeutic protocols for GBM. A label-free, mass spectrometry–based quantitative proteomics has been developed to identify and characterize proteins that are differentially expressed in GBM to gain a better understanding of the interactions and functions that lead to the pathological state focusing on the extracellular matrix (ECM). The main challenge in GBM research has been to identify novel molecular therapeutic targets and accurate diagnostic/prognostic biomarkers. To better investigate the GBM secretome upon in vitro treatment with histone deacetylase inhibitor (iHDAC), we employed a high-throughput label-free methodology of protein identification and quantification based on mass spectrometry followed by in silico studies. Our analysis revealed significant changes in the ECM protein profile, particularly those associated with the angiogenic matrisome. Proteins such as decorin, ADAM10, ADAM12, and ADAM15 were differentially regulated upon in silico analysis. In contrast, key angiogenesis markers such as VEGF and ECM proteins like fibronectin and integrins did not display significant changes. These results suggest that iHDAC inhibitors may modulate or suppress tumor behavior growth by targeting ECM proteins’ secretion rather than directly inhibiting angiogenesis.
Key features
• Analysis of the secretome of U87MG glioblastoma cells.
• Studies of mass spectrometry designed to modulate GBM biology and behavior focused on histone deacetylase inhibitors (iHDAC).
• Mass spectrometry was developed to identify and characterize proteins that are differentially expressed in GBM.
Keywords: Epigenetics Histone Deacetylase Glioblastoma Mass spectrometry Secretome
Graphical overview
Background
The identification of novel molecular therapeutic targets and accurate diagnostic/prognostic biomarkers has been a major challenge in glioblastoma (GBM) research. Proteins have the potential to be used as diagnostic and prognostic biomarkers in patients with brain tumors. They can be detected in glioblastoma cells and in liquid matrices such as blood and its derivatives, cerebrospinal fluid (CSF), and urine [1]. Currently, the main approach in the search for tumor markers is the study of proteomic profiling, along with the study of gene expression [2]. In the repertoire of tumor-associated proteins, a great variety of functional proteins, peptides, and other biomolecules are secreted or released by tumor cells to promote abnormal cell growth, invasion, and metastasis. Some of these proteins include growth factors, chemokines, and angiogenic factors, resulting in ECM remodeling, modulation of cellular signaling, and inflammatory responses in the tumor microenvironment. These proteins belong to pathways classically related to alterations in the tumor microenvironment and, for this reason, might be eligible as biomarkers for tracking tumor growth or relapse in non-invasive strategies upon surgery, chemo, and radiotherapy [3,4]. In this sense, the tumor secretome may reflect tumor behavior and predict patient management. In this work, we studied the alterations in the GBM secretome upon the treatment of U87-MG cells with a histone deacetylase inhibitor (iHDAC). In fact, preclinical studies have demonstrated the efficiency of different iHDACs as antitumor agents, especially when associated with other therapies, including chemotherapy and radiotherapy [5]. The main mechanisms involved in iHDACs efficiency as anti-tumor drugs are related to the reduction in the expression of genes involved in DNA repair, mitotic spindle formation, homologous chromosome segregation, and positive modulation of apoptosis [6].
The translation of iHDAC inhibitors into clinical practice requires a deeper understanding of their mechanism of action. To advance the clinical application of iHDAC inhibitors, it is essential to understand their impact on the tumor mechanisms that shape their microenvironment. By combining epigenetic and biochemical profiling, we can uncover the link between iHDACs and secreted molecular factors that drive tumor growth and potentially lead to new therapeutic strategies [7].
To understand how iHDAC inhibitors influence glioblastoma (GBM) secretome, we examined the total set of proteins secreted by iHDAC-treated GBM cells using mass spectrometry. Our analysis revealed significant changes in the extracellular matrix (ECM) protein profile, particularly those associated with the angiogenic matrisome. Proteins such as decorin, ADAM10, ADAM12, and ADAM15 were differentially regulated upon in silico analysis. In contrast, key angiogenesis markers such as VEGF and the ECM proteins fibronectin and integrins did not display significant changes. These results indicate that iHDAC inhibitors may modulate tumor behavior by targeting the ECM proteins’ secretion, rather than directly inhibiting angiogenesis [7].
Here, we describe a step-by-step protocol for our GBM cell line secretome analysis protocol, further categorization of tumor-secreted proteins, and the characterization of pathways in which these proteins play already known roles.
Materials and reagents
Biological materials
1. U87MG glioblastoma cell line (Banco de Células do Rio de Janeiro, BCRJ Code: 0241)
Reagents
1. Dulbecco's modified Eagle's medium (DMEM) low glucose (Sigma-Aldrich, catalog number: D5523)
2. Fetal bovine serum (FBS) (LGC Biotecnologia, catalog number: 10BioPlus-500)
3. Penicillin and streptomycin (PS) (Sigma-Aldrich, catalog number: P4333)
4. DMEM low-glucose without red phenol (Sigma-Aldrich, catalog number: D2902)
5. Trichostatin A (TSA) (Sigma-Aldrich, catalog number: T1952)
6. Sodium chloride (NaCl) (Sigma-Aldrich, catalog number: S3014)
7. Magnesium chloride (MgCl2·2H2O) (Sigma-Aldrich, catalog number: M8266)
8. Calcium chloride dihydrate (CaCl2·2H2O) (Sigma-Aldrich, catalog number: 223506)
9. Potassium chloride (KCl) (Sigma-Aldrich, catalog number: P3911)
10. Sodium phosphate dibasic (Na2HPO4) (Sigma-Aldrich, catalog number: S9763)
11. Dimethyl sulfoxide (DMSO) (Sigma-Aldrich, catalog number: D2650)
12. HaltTM protease inhibitor cocktail 100× (Thermo Fisher, catalog number: 78429)
13. QubitTM Protein and Protein Broad Range (BR) Assay kits (Invitrogen, catalog number: Q33211)
14. Urea (Sigma-Aldrich, catalog number: U4884)
15. Thiourea (Sigma-Aldrich, catalog number: T8656)
16. HEPES [4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid] (Sigma-Aldrich, catalog number: 5310-OP)
17. DL-Dithiothreitol (DTT) powder (Sigma-Aldrich, catalog number: 43819-5g)
18. Iodoacetamide (IAA) powder (Sigma, catalog number: I1149-25g)
19. Sequencing-grade modified trypsin (Promega, catalog number: V5111), see the product information
20. Formic acid LC–MS grade (Thermo Fisher Scientific, catalog number: 28905)
21. Acetonitrile (ACN) (Sigma-Aldrich, catalog number: 900667)
Solutions
1. Phosphate buffer saline (PBS) (see Recipes)
2. NaCl 2 M solution (see Recipes)
3. Urea lysis buffer (see Recipes)
4. DTT solution (see Recipes)
5. IAA solution (see Recipes)
6. Solvent A (ACN 5% and formic acid 0.1%) (see Recipes)
7. Solvent B (ACN 95% and formic acid 0.1%) (see Recipes)
Recipes
1. Phosphate buffer saline 10× (PBS)
Note: Dissolve all reagents in 800 mL of H2O. Adjust the pH to 7.4 (or 7.2, if required) with HCl, and then add H2O to 1,000 mL.
Reagent Final concentration (10×) Quantity or Volume
NaCl 137 mM 80 g
KCl 2.7 mM 2 g
Na2HPO4 10 mM 14.4 g
KH2PO4 1.8 mM 2.4 g
CaCl2·2H2O 1 mM 1.33 g
MgCl·2H2O 0.5 mM 1.0 g
H2O MilliQ n/a 800 mL
Total n/a 1,000 mL
2. NaCl 2 M solution
Note: Dissolve 116.9 g of NaCl in approximately 800 mL of H2O MilliQ, then add more water until a final volume of 1,000 mL.
Reagent Final concentration Amount
NaCl 2 mol/L 116.9 g
H2O MilliQ n/a 800 mL
Total n/a 1,000 mL
3. Urea lysis buffer
Note 1: Dissolving urea is an endothermic reaction. The preparation can be facilitated by placing a stir bar in the beaker and using a warm water bath on a stir plate. Attention, this water should not be too hot! To prevent urea from precipitating, the buffer should be used at room temperature.
Note 2: The urea lysis buffer should be prepared before each experiment. Aliquots can be stored in the -80 °C freezer for up to 6 months.
Reagent Final concentration Amount
Urea 7 mol/L 42.04 g
Thiourea 2 mol/L 15.22 g
HEPES 0.2 mol/L 4.76 g
H2O MilliQ n/a 80 mL
Total n/a 100 mL
4. DTT solution
Note: Dissolve 5 g of DTT in 324.14 mL of distilled water. After weighing the powder, store at -20 °C for up to one year. Thaw one aliquot for each experiment.
Reagent Final concentration Amount
DTT 100 mM 5 g
H2O MilliQ n/a 324.14 mL
5. Iodoacetamide (IAA) solution
Note: Dissolve 25 g of iodoacetamide in 338 mL of distilled water. After weighing the powder, store in the dark and add water only immediately before use. The iodoacetamide solution should be prepared just before use in each experiment.
Reagent Final concentration Amount
Iodoacetamide 400 mM 25 g
H2O MilliQ n/a 338 mL
6. Solvent A
Reagent Final concentration Amount
Acetonitrile 5% 0.5 mL
Formic acid 0.1% 0.01 mL
H2O MilliQ n/a 10 mL
7. Solvent B
Reagent Final concentration Amount
Acetonitrile 95% 9.5 mL
Formic acid 0.1% 0.01 mL
H2O MilliQ n/a 10 mL
Laboratory supplies
1. Pipettes (Eppendorf)
2. Tips (Eppendorf)
3. 50 mL Falcon centrifuge tubes, polypropylene, sterile (Corning, catalog number: 352070)
4. Microcentrifuge tube (Corning, Axygen, catalog number: MCT-150-C)
5. Becker
6. Water baths
7. Analytical Balance
8. 150 cm2 U-shaped canted neck cell culture flask with plug seal cap (Corning, catalog number: 430823)
9. Amicon Ultra-15 centrifugal filter unit (Meck Millipore, catalog number: UFC900324)
10. Pierce® C18 tips (87784 Pierce C18 Tips, 100 μL, 96 tips)
Equipment
1. Centrifuge
2. Centrifuge Heraeus Megafuge 8R (Thermo Fisher Scientific, catalog number: 75007214)
3. Sorvall centrifuge (Thermo Fisher Scientific, model: Lynx 4000)
4. CO2 incubator for cell culture (Thermo Fisher Scientific, model: 310 Series)
5. Speedvac (Thermo Fisher Scientific, model: SC210P1-15)
6. Thermomixer R with 1.5 mL block (Eppendorf, catalog number: Z605271)
7. -20 °C freezer (Thermo Fisher Scientific)
8. -80 °C freezer (Thermo Fisher Scientific)
9. Qubit protein assay kit, 500 assays (Thermo Fisher, catalog number: Q33238)
10. NanoLC (liquid chromatography) (Thermo Fisher Scientific, model: Easy1000)
11. Mass spectrometer (Thermo Fisher Scientific, model: Q ExactiveTM Plus)
12. Fused silica tubing (IDEX Health and Science, catalog number: FS-110)
13. PicoTipTM Emitter PicoFritTM SELF/P Capillary column (New Objective, catalog number: PF360-75-15-N-5)
14. Reprosil-Gold C18 (Dr. Maisch, catalog number: r33.9g-3g)
15. Reprosil-Pur C18 (Dr. Maisch, catalog number: r23.aq-2g)
Software and datasets
1. Software Xcalibur, Thermo Fisher Scientific, 2.2, 2011
2. Proteome Discoverer (PD), Thermo Fisher Scientific, 2.1, 2015
3. Uniprot database, 2017. Available in: https://www.uniprot.org/
4. All data have been deposited in ProteomeXchange Consortium via the PRIDE. Available at https://www.ebi.ac.uk/pride/ with the dataset identifier PXD022982 at PRIDE username: [email protected] password: OBpIGRa2.
5. Microsoft Excel (Microsoft) or equivalent spreadsheet tool
Procedure
A. Preparation of conditioned medium–secretome
1. Seed U87MG GBM cells at a density of 7 × 105 cells on 150 cm2 flasks with DMEM supplemented with 10% FBS and 0.1 mg/mL PS until they reach 90% confluence.
2. Maintain cultures at 37 °C with 5% CO2.
3. Wash cell cultures once with cold PBS, once with cold 2 M NaCl solution, and then twice with cold PBS to remove residues of FBS.
4. Add 50 mL of DMEM without phenol red and FBS to each 150 cm2 flask.
5. Add DMEM, low glucose, without phenol red and FBS, supplemented with trichostatin A at a concentration of 100 nM to the group under HDAC activity inhibition (here referred to as iHDAC).
6. Add the same media supplemented only with the vehicle dimethyl sulfoxide (DMSO) to the control group.
7. Carry out treatments for 72 h at 37 °C with 5% CO2.
B. Dialysis of the conditioned medium–secretome
1. Collect the conditioned media after 72 h of treatment and place in a 50 mL Falcon tube. After centrifugation at 1,200 rpm (258× g) for 10 min, collect the secretome to remove cells and debris. For secretome concentration, use AMICON ULTRA 4 3kD 50 mL tubes. Due to the limitation of volume in the AMICON tube, add only 5 mL of secretome at a time to the top of the tube and centrifuge at 5,000 rpm (3,075× g) for 30 min until the total secretome is centrifuged. At the end, concentrate the secretome until it reaches the final volume of 250 μL.
2. Add protease inhibitor Halt 1× and perform protein quantification using the Qubit Protein Assay Kit method according to the manufacturer's instructions. As a blank, use DMEM medium without phenol red.
C. Protein digestion and peptide purification
1. Dilute 100 μg of total protein in 20 μL of 7 M urea and 2 M thiourea in 0.2 M HEPES.
2. Add DTT at a final concentration of 10 mM, followed by incubation for 1 h at 30 °C in the thermomixer without stirring.
3. After incubation, add IAA at a final concentration of 40 mM and then incubate in the dark for 30 min at room temperature.
4. Dilute the solution ten-fold with standard LC–MS deionized water and add trypsin at a final concentration of 1:50 trypsin/protein (w/w). Incubate the mixture overnight at 37 °C in the thermomixer under 900 rpm rotation. (One vial of 20 μg of trypsin was diluted in 200 μL of 0.1 M acetic acid solution; this solution is provided with the trypsin.)
5. Add 1% formic acid to stop trypsin digestion and desalt the peptides using C18 microcolumns (ZipTip).
6. Transfer the purified peptides to injection vials and send to NanoLC-MS/MS analysis.
D. Analysis by NanoLC-MS/MS
1. Analyze the peptides using an Easy1000 nanoLC system coupled to a Q ExactiveTM Plus mass spectrometer.
2. Measure the concentration of peptides using the QubitTM Protein Assay kit.
3. For each sample, apply a volume of 4 μL (1 μg of peptides) to a Trap column with 200 μm of internal diameter and 2 cm long packed with Reprosil-Pur C18 with pores of 200 Å and 5 μm particle size (made at the laboratory).
4. Elute peptides in an analytical column with 75 μm diameter and 25 cm length packed with Reprosil-Gold C18 with pores of 300 Å of 3 μm granulometry (made at the laboratory).
5. For the peptide separation, use a gradient of 95% solvent A (5% ACN and 0.1% formic acid) to 20% of solvent B (95% ACN and 0.1% formic acid) for 120 min, 20%–40% solvent B for 40 min, and 40%–95% solvent B for 7 min, followed by 95% solvent B for 13 min.
6. Afterward, rebalance the column with solvent A.
7. Set the Orbitrap mass spectrometer, controlled by the Xcalibur 2.2 software, to operate with the data-dependent acquisition (DDA) mode.
8. Acquire the spectrum of MS1 with a resolution of 70,000 to 200 m/z (mass/charge).
9. Perform the reading of the MS1 spectrum using 10e6 ions (AGC) and 50 ms of Maximum IT.
10. The reading spectrum comprised ions with 375–2,000 m/z. The 15 most intense ions were fragmented and then subjected to MS2 acquisition using induced collision dissociation (HCD) and a range of 200–2,000 m/z.
11. Apply MS2 resolution at 17,500 to 200 m/z, AGC of 10e5 ions, maximum IT of 100 ms, 2 m/z ion isolation window, normalized collision energy (NCE) of 30, and dynamic exclusion time 45 s.
12. Reject peptides with undetermined charges and +1.
13. Inject peptide fractions three times to obtain three technical replicates for each one of the two biological samples of each experimental group.
E. Data analysis
1. The peptides were identified using the Sequest HT search engine.
2. Process raw files using the Proteome Discoverer (PD) 2.1 software.
3. The Uniprot database used was the Homo sapiens reference proteome (June 2017). Common contaminants of the Global Proteome machine (GPM) were used to exclude putative contaminants; after this first analysis, 42,227 protein entries were obtained.
4. The PD Sequest HT node parameters applied were: full-tryptic search space with two missed cleavages allowed, precursor mass tolerance of 10 ppm, and fragment mass tolerance of 0.05 Da.
5. Carbamidomethylation of cysteine (C) was included as a constant modification, and methionine (M) oxidation and N-terminal acetylation of proteins were included as dynamic modifications.
6. The Percolator node in Proteome Discovery 2.1 (PD 2.1) was used to estimate the false discovery rate (FDR) of <1% and protein inference using maximum parsimony.
7. A cutoff was set to accept a 1% FDR at the protein and peptide levels.
8. Protein quantification was performed using the precursor ions area detector node, with an average of up to three most intense peptides used for protein quantification, which were considered unique peptides.
9. The table of identified proteins was exported for bioinformatics and statistical studies.
10. Mass spectrometry proteomics data were submitted to the ProteomeXchange Consortium via PRIDE.
F. Statistical rationale
1. The effect of iHDAC treatment on the U87-MG GBM secretome analysis was evaluated after 72 h of treatment with 100 nM trichostatin A or DMSO as control.
2. The iHDAC and control secretomes were obtained from two biological replicates of each condition, and technical triplicates of LC–MS/MS runs were conducted for each experimental sample.
3. To check the reproducibility of the replicates, Pearson correlation was used to correlate the three technical runs of each biological sample. Next, the common proteins identified in the two biological replicates of iHDAC (TSA1 and TSA2) or in the two biological replicates of the control group (DMSO1 and DMSO2) were taken as the identified proteins in each experimental group and used to further compare the differentially secreted proteins in iHDAC and DMSO secretome.
4. Statistical analysis was carried out using Perseus v 1.6.10.50 and R. Protein area values were converted to log2 scale and normalized by subtracting the median of the sample distribution. Proteins with <50% valid values in each group (iHDAC and DMSO) were removed from the analysis.
5. The remaining proteins were subjected to missing value imputation using the default parameters (width 0.3, downshift 1.8) of the Perseus Imputation tool. GraphPad Prism (v6.0, La Jolla, CA) was used for ordinary one-way or two-way ANOVA analysis. If the ANOVA produced a significant result, post-hoc pairwise comparisons were tested for significance in which the value was adjusted (Padj < 0.05) by Tukey’s method for multiple comparisons inside each group.
6. Results were presented as mean ± SEM and statistical relevance was defined as p < 0.05.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
• Menezes et al. [7]. Epigenetic Mechanisms Histone Deacetylase–Dependent Regulate the Glioblastoma Angiogenic Matrisome and Disrupt Endothelial Cell Behavior In Vitro. Molecular & Cellular Proteomics, Volume 23, Issue 3, 100722.
Acknowledgments
We thank the Unidade de Espectrometria de Massas e Proteômica (UEMP) from the Universidade Federal do Rio de Janeiro for the use of the facilities for samples preparation, Grasiella Ventura Matioszek for Confocal support, CAPES (fellowship for M.A. and J.G.), FAPERJ and Fundação do Câncer Ary Frauzino.
Funding and additional information: C.K. was supported by FAPERJ/-26/010.001760/2019 and CNPq 315751/2021-5.
Competing interests
The authors declare that they have no conflicts of interest with the contents of this article.
References
Hosseini, A., Ashraf, H., Rahimi, F., Alipourfard, I., Alivirdiloo, V., Hashemi, B., Yazdani, Y., Ghazi, F., Eslami, M., Ameri Shah Reza, M., et al. (2023). Recent advances in the detection of glioblastoma, from imaging-based methods to proteomics and biosensors: A narrative review. Cancer Cell Int. 23(1): 98.
Silantyev, A., Falzone, L., Libra, M., Gurina, O., Kardashova, K., Nikolouzakis, T., Nosyrev, A., Sutton, C., Mitsias, P., Tsatsakis, A., et al. (2019). Current and Future Trends on Diagnosis and Prognosis of Glioblastoma: From Molecular Biology to Proteomics. Cells. 8(8): 863.
Bertucci, F. and Goncalves, A. (2008). Clinical Proteomics and Breast Cancer: Strategies For Diagnostic and Therapeutic Biomarker Discovery. Future Oncol. 4(2): 271–287.
Sanders, M. E., Dias, E. C., Xu, B. J., Mobley, J. A., Billheimer, D., Roder, H., Grigorieva, J., Dowsett, M., Arteaga, C. L., Caprioli, R. M., et al. (2008). Differentiating Proteomic Biomarkers in Breast Cancer by Laser Capture Microdissection and MALDI MS. J Proteome Res. 7(4): 1500–1507.
Shabason, J. E., Tofilon, P. J. and Camphausen, K. (2011).Grand rounds at the National Institutes of Health: HDAC inhibitors as radiation modifiers, from bench to clinic. J Cell Mol Med. 15(12): 2735–2744.
Cornago, M., Garcia-Alberich, C., Blasco-Angulo, N., Vall-llaura, N., Nager, M., Herreros, J., Comella, J. X., Sanchis, D. and Llovera, M. (2014). Histone deacetylase inhibitors promote glioma cell death by G2 checkpoint abrogation leading to mitotic catastrophe. Cell Death Dis. 5(10): e1435.
Menezes, A., Julião, G., Mariath, F., Ferreira, A. L., Oliveira-Nunes, M. C., Gallucci, L., Evaristo, J. A. M., Nogueira, F. C. S., Pereira, D. d. A., Carneiro, K., et al. (2024). Epigenetic Mechanisms Histone Deacetylase–Dependent Regulate the Glioblastoma Angiogenic Matrisome and Disrupt Endothelial Cell Behavior In Vitro. Mol Cell Proteomics. 23(3): 100722.
Article Information
Publication history
Received: Sep 7, 2024
Accepted: Dec 17, 2024
Available online: Jan 14, 2025
Published: Feb 5, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Category
Cancer Biology > Angiogenesis > Tumor microenvironment
Systems Biology > Proteomics > Secretome
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5,198 | https://bio-protocol.org/en/bpdetail?id=5198&type=0 | # Bio-Protocol Content
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Flood Inoculation of Fusarium eumartii in Tomato Seedlings: Method for Evaluating the Infectivity of Pathogen Spores
MT María Cecilia Terrile
FM Florencia Anabel Mesas
MP María Elisa Picco
MS María Florencia Salcedo
AM Andrea Yamila Mansilla
Published: Vol 15, Iss 3, Feb 5, 2025
DOI: 10.21769/BioProtoc.5198 Views: 42
Reviewed by: Noelia ForesiMalgorzata Lichocka Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Pest management science May 2014
Abstract
The Fusarium genus includes various fungi of great significance in agriculture. Fusarium solani f. sp. eumartii (F. eumartii), traditionally known as a potato pathogen, has also been identified as a cause of disease in tomatoes. This protocol provides a detailed, efficient, and robust flood-inoculation method for assessing F. eumartii infection of young tomato seedlings grown on MS medium plates. It includes the evaluation of the lesion area and the quantification of the remaining fungal inoculum in tomato seedlings. In summary, the straightforwardness and efficiency of this bioassay make it a powerful quantitative tool for selecting fungicidal compounds or defense response inducers in tomato plants, offering a promising approach with significant potential for preventing fungal diseases in crops.
Key features
• Accelerated fungal infection method in tomato seedlings, shortening overall experimental time.
• Allows simultaneous evaluation of fungal infectivity and quantitative remaining inoculum.
• Easily adaptable for screening fungicides and defense inducers in various plant–pathogen interactions.
Keywords: Fusarium Tomato Bioassay Flood inoculation Pathogenicity
Graphical overview
Graphical representation of the protocol for assessing the infectivity of Fusarium solani f. sp. eumartii in tomato seedlings using a flood-inoculation method. The diagram outlines the main steps: i) preparation of the inoculum from fungal mycelium, ii) germination and growth of tomato seedlings, iii) inoculation of seedlings with the spore suspension, iv) infection assessment by measuring lesion areas and remaining fungal inoculum, and v) data analysis. Arrows indicate the sequence of the experimental procedure.
Background
Numerous species within the Fusarium genus, such as Fusarium oxysporum and Fusarium solani, are responsible for significant agricultural losses worldwide, both before and after harvest [1]. Fusarium solani f. sp. eumartii (F. eumartii) is a significant post-harvest disease affecting potato tubers, leading to reddish-brown mottling between leaf veins and dry rot [2]. While traditionally recognized as a potato pathogen (Solanum tuberosum), F. eumartii has also been identified in tomato (Solanum lycopersicum) plants. Isolates from these plant species have also shown pathogenic effects in other members of the Solanaceae family [3]. Different methods of infecting tomato plants with Fusarium fungi have been reported in the literature. In this protocol, we describe a simple, reliable, and reproducible way to assess the infection of tomato seedlings with this pathogen. This method was adapted from a seedling-based assay initially developed by Uppalapati et al. [4] for Pseudomonas syringae pv. tomato, demonstrating its versatility and effectiveness in studying plant-pathogen interactions. Moreover, this approach allows a practical screening of different compounds that can act with fungicides, as described in the work of Terrile et al. [5].
Materials and reagents
Biological materials
1. Fusarium solani f. sp. eumartii (F. eumartii) isolated 3122 (Estación Experimental Agropecuaria -EEA-INTA, Balcarce, Argentina; Radtke and Escande, 1973)
2. Tomato (Solanum lycopersicum cv. Platense) was obtained from a commercial supply
Reagents
1. Agar (BD-DIFCO, catalog number: 214530)
2. Ampicillin (Amp) (GenBiotech, catalog number: A-0104-5)
3. Commercial bleach (sodium hypochlorite <5%)
4. Ethanol (Biopack, catalog number: 2000165400)
5. Murashige and Skoog (MS) medium (Sigma, catalog number: M5524)
6. KOH (Sigma, catalog number: 221473)
7. Potato dextrose agar (PDA) (OXOID, catalog number: CM0139)
8. Silwet L77 (Momentive, catalog number: A0300026)
9. Sterile distilled water
10. Triton X-100 (Biopack, catalog number: 2000200308)
Solutions
1. MS medium, 1 L (see Recipes)
2. PDA medium, 1 L (see Recipes)
3. 30% bleach solution (see Recipes)
4. 70% ethanol (see Recipes)
Recipes
1. MS medium (1 L)
4.4 g of MS basal medium
8 g of agar
Adjust to pH 5.7 with 1 M KOH and autoclave at 121 °C for 15 min
2. PDA medium (1 L)
39 g of PDA
Autoclave at 121 °C for 15 min
3. 30% bleach solution
Add 3 mL of bleach into 7 mL of distilled water and 0.04% Triton X-100
4. 70% ethanol
Add 73 mL of ethanol 96% into 27 mL of distilled water.
Laboratory supplies
1. Forceps
2. Conical tubes (15 and 50 mL) (Deltalab, catalog number: 429942 and 429931)
3. Household aluminum foil for multiple uses
4. 1.5 mL microcentrifuge tubes (Deltalab, catalog number: 4092.4N)
5. Neubauer chamber (Marienfeld, catalog number: 0610101)
6. Paper towels
7. Parafilm (Amcor, catalog number: PM-996)
8. Petri dishes 90 × 14 mm (Deltalab, catalog number: 200200)
9. 0.5 mm diameter punching tool
10. 5 mL pipette tips (Deltalab, catalog number: 900038.C)
11. 1 mL pipette tips (Deltalab, catalog number: 200012)
12. 2–20 μL pipette tips (Deltalab, catalog number: 200016B)
13. Micro-pestle (Axygen, catalog number: AXYPES15BSI)
Equipment
1. Cultivation room at 25 °C under 250 μmol photons m-2 s-1 with 70% ± 5% relative humidity and 16:8 h light/dark cycle
2. Cold room at 4 °C
3. Laminar air flow equipment (CASIBA, model: 3101-FLV-2)
4. Pipettes: PIPETMAN P20L, P200, P1000, P5000 (Gilson, model: F144056M, F144058M, F144059M and F144066)
5. Vortex (VELP Scientifica, model: ZX-3)
6. Stereomicroscope (Nikon, model: SMZ800)
7. Digital camera (Canon, model: DS126431)
Software and datasets
1. ImageJ software (NIH, https://imagej.net/)
2. Graphpad Prism version 5.01 software for Windows (GraphPad Software, La Jolla, California, USA, 2007)
Procedure
A. Fungal growth and inoculum preparation
1. Critical step: Prepare PDA plates supplemented with Amp (100 μg/mL) to prevent bacterial contamination during the growth of F. eumartii while maintaining the viability and functionality of fungi.
2. Cut a 0.5 mm PDA disc with sterile punching tools from plates previously colonized with F. eumartii mycelium.
3. Place the disc in the center of the freshly prepared plate, with the mycelium toward the culture medium (Figure 1A).
4. Incubate the plates in the cultivation room at 25 °C for 6–7 days in darkness. At this time, all the plates are covered with fungal mycelium (Figure 1B).
5. Store the plates at 4 °C in darkness.
6. To prepare inoculum (spore suspension) for the bioassay, scrape the mycelium superficially with a sterile spatula to obtain spores and submerge in a 1.5 mL microcentrifuge tube with 1 mL of sterile distilled water (Figure 1C).
7. Count the spores in a Neubauer chamber.
8. Dilute the original spore suspension to obtain a concentration of 107 spores/mL.
Note: If the action of a fungicidal compound is to be evaluated, treatment and incubation of the spores can be performed at this point before plant infection.
9. Add Silwet L77 to achieve a final concentration of 0.025% (v/v) and mix thoroughly.
Note: Use the fungal suspensions immediately after preparation.
Figure 1. Fungal growth and inoculum preparation. A. Inoculation of freshly prepared PDA plate with fungus-colonized PDA disc. B. 7-day F. eumartii mycelia growth on PDA plate. C. Scraping of mycelium to obtain a spore solution to inoculate plants.
B. Plant growth
1. Seed sterilization
a. Incubate 500 tomato seeds (approximately 1 g) in 10 mL of 70% ethanol for 2 min in a 15 mL conical tube.
b. Discard the ethanol solution by pipetting. Caution: Rinse the seeds twice with sterile distilled water to avoid ethanol residues.
c. Incubate the seeds with 10 mL of 30% bleach solution for 15 min with gentle shaking (500–1,000 rpm).
d. Wash the seeds thoroughly with 10 mL of sterile distilled water three times. Caution: Ensure complete removal of bleach residues to prevent seed damage or interference with germination.
Pause point: The seeds can be kept in sterile distilled water at 4 °C for a few hours before sowing on the MS plates.
2. Seed germination
a. Sow the seeds with sterile forceps in Petri dishes with MS medium (20–30 seeds per plate; Figure 2A).
Note: Prepare at least three plates per treatment for quantitative analysis.
b. Incubate the plates horizontally in the cultivation room at 25 °C for 5–6 days until cotyledons emerge.
Figure 2. Plant growth and F. eumartii flood inoculation of tomato seedlings. A. Sowing of seed on MS plates. B. Transferring 5-day-old healthy seedlings to fresh MS medium plate. C. Inoculation with spore suspension into cotyledons.
C. In planta bioassay
1. Carefully transfer the healthy seedlings with fully emerged cotyledons to fresh MS medium plates (five seedlings per plate) using sterile forceps (Figure 2B). Ensure that the cotyledons and roots are handled gently to avoid physical damage, which could compromise seedling health or affect infection outcomes.
Note: If desired, transfer the seedlings to MS medium plates supplemented with the treatments to be tested as fungicides.
2. Add 5 mL of the spore suspension to each Petri dish, ensuring that the tomato cotyledons are thoroughly moistened. Incubate for 10–15 min, occasionally mixing the solution to maintain even coverage (Figure 2C). As a control, add 5 mL of a solution containing sterile distilled water and Silwet L77 (at a final concentration of 0.025% v/v) to a separate set of Petri dishes with tomato seedlings.
3. Discard the inoculum suspension and seal the plates with parafilm.
4. Incubate the plates in the cultivation room at 25 °C for 5–6 days until disease symptoms appear.
Caution: To ensure accuracy and reliability, independent sets of cotyledons are used to quantify the lesion area and measure the remaining fungal inoculum. Cotyledons selected for lesion area quantification remain undisturbed for imaging and measurement, while those allocated for inoculum analysis are processed separately to minimize cross-contamination and variability between analyses.
D. Quantification of tomato lesion area
1. Take a photograph of each tomato cotyledon under a stereomicroscope to capture detailed images of the lesion areas (Figure 3A).
Note: To standardize image quality, pictures of tomato cotyledons were taken under consistent lighting conditions using a stereomicroscope equipped with a ring light (intensity set to 75%) to minimize shadows. A digital camera (Canon DS126431) was set to manual mode. Images were saved in JPEG format at maximum resolution.
Figure 3. Visual infection symptoms and quantification of remaining fungal inoculum. A. Symptoms of infection in cotyledons (left, pronounced symptoms; right, mild symptoms). B. Homogenizing to effectively recover the remaining inoculum. C. Counting CFU after serial dilutions.
2. Measure the lesion area in the photographed cotyledons using the image-processing software ImageJ. Convert the images to an 8-bit grayscale format to enhance contrast and facilitate measurements. To measure one lesion area (Figure 3A, left cotyledon), use the polygon selection tool to delineate the total area of the cotyledon and select the measure tool (Ctrl+M) to obtain the area value, which will be considered 100%. Repeat the previous sequence by selecting the infected area. Record the area value for the infected region and calculate the lesion area percentage as follows: (infected area/total area) × 100.
3. To analyze multiple lesion areas (Figure 3A, right cotyledon), use the polygon selection tool to delineate each infected region on the cotyledon. After selecting each region, add it to the ROI manager by pressing "Add [t]" in the ROI manager window. Repeat this process for all visible lesions. Then, select the measure tool to obtain the cumulative area of all regions and calculate the lesion area percentage as follows: (cumulative infected area/total area) × 100.
E. Measurement of residual fungal inoculum
1. Remove the cotyledons from the seedlings of each plate, place them in a 15 mL conical tube, and add 10 mL of 70% ethanol for 1 min to eliminate epiphytic bacteria before sampling. Rinse the plant tissue gently with sterile distilled water.
2. Dry the tissue with regular paper towels. Collect approximately 50 mg of the set of cotyledons and place them in a 1.5 mL microcentrifuge tube.
Note: Cotyledons from two independent replicate plants are pooled for a single tissue sample. Three or more samples are needed for each treatment.
3. Add 500 μL of cold sterile distilled water to the tube and homogenize with a plastic micro-pestle by hand until pieces of tissue are no longer visible (Figure 3B). Critical: Ensure the tissue is thoroughly homogenized to prevent discrepancies in CFU counts.
4. Vortex the homogenates for 5 s.
5. Serially dilute the sample in sterile distilled water.
Note: The number of serial dilutions required to obtain countable colonies should be determined empirically for each sample, but dilutions up to 10–7 are usually sufficient.
6. Plate the dilutions in PDA plates supplemented with 100 μg/mL Amp and incubate at 25 °C for 2 days in darkness. Count the CFU using appropriately diluted samples (Figure 3C).
Data analysis
Analyze the data set with Student’s t-test for comparing two averages of fungal growth or ANOVA with post-hoc Tukey’s multiple range test to compare more than two averages of fungal growth.
Validation of protocol
This protocol has been used and validated in the following research article:
Terrile et al. [5]. Nitric oxide–mediated cell death is triggered by chitosan in Fusarium eumartii spores. Pest management science.
The key findings presented in this reference work are summarized as follows:
The validation results highlight the dual effectiveness of chitosan against Fusarium eumartii in tomato seedlings through direct fungicidal action and induced plant defenses. In seedlings inoculated with chitosan-treated spores, necrotic symptoms were reduced by approximately 90%, and residual fungal inoculum decreased by 83% compared to untreated controls. Additionally, pre-treating seedlings with 10 μg/mL of chitosan for 4 days significantly reduced both lesion area and remaining fungal inoculum following spore inoculation. These findings confirm that chitosan not only inhibits fungal growth but also acts as an elicitor of plant defense mechanisms, enhancing the plant's resistance to infection.
General notes and troubleshooting
General notes
The F. eumartii plates used to obtain spore solution should not be more than 3–4 weeks old because older plates may have reduced spore viability or altered fungal physiology/pathogenicity. This could impact the consistency and reliability of the inoculum, leading to variability in infection outcomes and potentially compromising the experiment's reproducibility and accuracy.
Troubleshooting
Problem 1: Absence of disease symptoms on tomato seedlings after 5–6 days post-inoculation.
Possible causes: The inoculum may not be infective, the inoculation may not have been efficient, or the environmental conditions in the growth chamber may not be suitable for infection.
Solution: Ensure that the inoculum is applied uniformly to the cotyledons. If symptoms are not observed after 5–6 days, extend the incubation period and monitor the cultivation room parameters (temperature, humidity, and light). To verify the viability of the inoculum used for inoculation, incubate spore suspension on multi-well microscope slides for 16 h at 25 °C and 100% humidity in darkness. Check for germination under a microscope. Germination should exceed 80% for viability.
Problem 2: Inconsistent quantification of fungal population in inoculated plant tissue.
Possible causes: Variability in the effectiveness of tissue homogenization, inconsistencies in sample preparation methods, or unintentional contamination during handling.
Solution: Use a precise weight of plant tissue for each sample, ensuring uniformity across replicates and treatments. Homogenize the tissue thoroughly to achieve a consistent suspension. Perform dilutions consistently using a fixed volume of homogenized tissue extract for initial dilution and use calibrated pipettes to ensure accuracy in volumes during dilution steps. Make positive control, including a known concentration of fungal spores to validate the dilution and plating process. Additionally, perform parallel quantification of CFU from an uninfected tissue sample as a negative control to account for contamination.
Acknowledgments
This work was partially supported by grants from Agencia Nacional de Promoción de la Investigación, el Desarrollo y la Innovación (Agencia I+D+i, PICT2019-3002 and PICT Start Up 0023), Consejo Nacional de Investigaciones Científicas y Técnicas (CONICET, PIP 0237); Fundación Williams and Universidad Nacional de Mar del Plata (EXA1137/23).
Competing interests
The authors declare that they have no conflict of interest.
References
K. Chehri. (2011). Occurrence of Fusarium species associated with economically important agricultural crops in Iran. Afr J Microbiol Res. 5(24): e158. https://doi.org/10.5897/ajmr10.158
D’Ippólito, S., Martín, M. L., Salcedo, M. F., Atencio, H. M., Casalongué, C. A., Godoy, A. V. and Fiol, D. F. (2010). Transcriptome profiling of Fusarium solani f. sp. eumartii -infected potato tubers provides evidence of an inducible defense response. Physiol Mol Plant Pathol. 75: 3–12. https://doi.org/10.1016/j.pmpp.2010.09.002
Romberg, M. K. and Davis, R. M. (2007). Host Range and Phylogeny of Fusarium solani f. sp. eumartii from Potato and Tomato in California. Plant Dis. 91(5): 585–592. https://doi.org/10.1094/pdis-91-5-0585
Uppalapati, S. R., Ishiga, Y., Wangdi, T., Urbanczyk-Wochniak, E., Ishiga, T., Mysore, K. S. and Bender, C. L. (2008). Pathogenicity of Pseudomonas syringae pv. tomato on Tomato Seedlings: Phenotypic and Gene Expression Analyses of the Virulence Function of Coronatine. Mol Plant-Microbe Interact. 21(4): 383–395. https://doi.org/10.1094/mpmi-21-4-0383
Terrile, M. C., Mansilla, A. Y., Albertengo, L., Rodríguez, M. S. and Casalongué, C. A. (2014). Nitric‐oxide‐mediated cell death is triggered by chitosan in Fusarium eumartii spores. Pest Manage Sci. 71(5): 668–674. https://doi.org/10.1002/ps.3814
Article Information
Publication history
Received: Sep 25, 2024
Accepted: Dec 15, 2024
Available online: Jan 5, 2025
Published: Feb 5, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
How to cite
Category
Plant Science > Plant immunity > Host-microbe interactions
Microbiology > Microbe-host interactions > Fungus
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5,199 | https://bio-protocol.org/en/bpdetail?id=5199&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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Peer-reviewed
Identification of Neurons Containing Calcium-Permeable AMPA and Kainate Receptors Using Ca2+ Imaging
SG Sergei G. Gaidin
AK Artem M. Kosenkov
VZ Valery P. Zinchenko
BK Bakytzhan K. Kairat
AM Arailim E. Malibayeva
ST Sultan T. Tuleukhanov
Published: Vol 15, Iss 3, Feb 5, 2025
DOI: 10.21769/BioProtoc.5199 Views: 47
Reviewed by: Alessandro Didonna Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Frontiers in Synaptic Neuroscience Mar 2024
Abstract
Calcium-permeable AMPA receptors (CP-AMPARs) and kainate receptors (CP-KARs) play crucial roles in synaptic plasticity and are implicated in various neurological processes. Current methods for identifying neurons expressing these receptors, such as electrophysiological recordings and immunostaining, have limitations in throughput or inability to distinguish functional receptors. This protocol describes a novel approach for the vital identification of neurons containing CP-AMPARs and CP-KARs using calcium imaging. The method involves loading neurons with Fura-2 AM, a calcium-sensitive fluorescent probe, KCl application to identify all neurons, and further addition of specific AMPAR agonists (e.g., 5-fluorowillardiine) in the presence of voltage-gated calcium channel blockers and NMDAR/KAR antagonists to identify CP-AMPAR-containing neurons. CP-KAR-containing neurons are identified using domoic acid applications in the presence and absence of NASPM (a CP-AMPAR antagonist). This technique offers several advantages over existing methods, including the ability to assess large neuronal populations simultaneously, distinguish between different receptor types, and provide functional information about CP-AMPAR and CP-KAR expression in living neurons, making it a valuable tool for studying synaptic plasticity and neurological disorders.
Key features
• The described protocol allows vital identification of neurons containing calcium-permeable AMPA (CP-AMPARs) and kainate receptors (CP-KARs).
• This approach can be combined with other methods, such as electrophysiological recordings or immunostaining.
• The method is fast, reproducible, and allows non-invasive simultaneous identification of numerous CP-AMPAR-/CP-KAR-containing neurons.
• The described protocol can be used for pharmacological screening of different drugs, including neuroprotectors, or investigation of features of CP-AMPAR-/CP-KAR-containing neurons in health and disease.
Keywords: Calcium-permeable AMPA receptors Calcium-permeable kainate receptors GABAergic neurons Identification Calcium imaging Intracellular Ca2+ concentration
Background
Glutamate is a primary excitatory neurotransmitter in the mammalian nervous system that activates various types of glutamate receptors, including AMPA receptors (AMPARs). A subset of AMPARs, known as calcium-permeable AMPA receptors (CP-AMPARs), play a crucial role in synaptic plasticity mechanisms such as long-term potentiation (LTP) and depression (LTD) [1,2]. In addition to Na+ and K+ conductance, these receptors are permeable for Ca2+ ions due to either the absence of the GluA2 subunit or the presence of the unedited GluA2 subunits [3].
Although CP-AMPARs are involved in normal brain functioning, they are also associated with various pathologies [4], including neurodegenerative diseases such as Alzheimer’s [5,6] and Parkinson’s diseases [7,8]. CP-AMPAR surface expression is dynamically regulated in response to changes in synaptic activity: while a weak stimulus promotes CP-AMPAR incorporation into the plasma membrane, stronger stimulation favors calcium-impermeable AMPARs [4]. This dynamic regulation underlies their importance in both Hebbian and non-Hebbian forms of plasticity across different brain regions.
Despite CP-AMPARs' significance in synaptic processes, our understanding of their role and the neurons expressing them remains poorly studied. This knowledge gap is partly due to the challenges associated with identifying CP-AMPAR-containing neurons. Current methods for detecting these receptors have notable limitations. Electrophysiological techniques, such as current-voltage relationship analysis and the use of polyamine antagonists, can identify synapses with CP-AMPARs and study their functions in individual neurons [9]. However, these approaches are labor-intensive and limited in their ability to assess large neuronal populations simultaneously. Additionally, some antagonists used in these studies may also affect kainate receptors, potentially confounding results [10]. Immunostaining methods, typically using anti-GluA2 antibodies, allow the evaluation of CP-AMPAR expression across larger groups of neurons [11,12]. However, this approach cannot distinguish between edited and unedited GluA2 subunits, which is crucial as unedited GluA2-containing AMPARs are also calcium-permeable. While most GluA2 subunits in the adult brain are edited, thus resulting in the formation of calcium-impermeable AMPARs, editing levels can change under certain pathological conditions, potentially leading to inaccurate conclusions about CP-AMPAR expression.
To address these limitations, we have developed a novel protocol based on vital fluorescent calcium imaging. This method offers several advantages over existing techniques:
1. It enables the identification of neurons expressing both GluA2-lacking CP-AMPARs and those containing unedited GluA2 subunits.
2. The approach visualizes neurons with a significant number of CP-AMPARs sufficient to induce detectable somatic calcium influx.
3. It facilitates the evaluation of changes in CP-AMPAR expression at the population level, allowing for the study of various experimental manipulations in models of brain pathologies.
4. The technique is less labor-intensive than electrophysiological recordings and provides more functional information than immunostaining alone.
Our calcium imaging-based protocol for identifying CP-AMPAR-containing neurons offers a valuable tool for advancing research in synaptic plasticity, neuronal development, and neuropathology. By bridging the gap between single-cell electrophysiology and population-level immunostaining, this method provides a more comprehensive understanding of CP-AMPAR distribution and function in the nervous system. We believe that it will shed light on the roles of these receptors in both physiological and pathological processes, opening new avenues for research into neurological disorders and potential therapeutic interventions.
Materials and reagents
Biological materials
1. Postnatal (P0-2) Wistar male rats (branch of the M.M. Shemyakin and Yu.A. Ovchinnikov Institute of Bioorganic Chemistry of the Russian Academy of Sciences)
Reagents
1. Poly(ethyleneimine) solution 50% w/v in water (Sigma-Aldrich, catalog number: P3143); stock and working solutions are stored at 4 °C
2. (±)-Verapamil hydrochloride (Sigma-Aldrich, catalog number: V4629); powder is stored at 4 °C, whereas aliquots are stored at -20 °C
3. Penicillin–streptomycin 100× solution (Sigma-Aldrich, catalog number: P4333); aliquots stored at -20 °C
4. L-Glutamine (Sigma-Aldrich, catalog number: G85402); powder is stored at 4 °C, whereas aliquots are stored at -20 °C
5. (S)-5-Fluorowillardiine (Sigma-Aldrich, catalog number: F2417); powder and aliquots are stored at -20 °C
6. Neurobasal-A medium (Life Technologies, catalog number: 10888022); stored at 4 °C.
7. 50× B27 supplement (Life Technologies, catalog number: 17504044); aliquots of stock solution (we recommend using 1 mL aliquots) are stored at -20 °C
8. Trypsin 2.5% (Life Technologies, catalog number: 15090046); stock solution and working solution aliquots are stored at -20 °C
9. Fura-2 AM (Molecular Probes, catalog number: F1221); undissolved dye and aliquots of the stock solution are stored at -20 °C; the stock solution should be bubbled with argon (recommended) or nitrogen and tightly sealed before freezing
10. Bicuculline (Cayman Chemical, catalog number: 11727); the powder is stored at 4 °C, whereas aliquots are stored at -20 °C
11. UBP310 (Tocris Bioscience, catalog number: 3621); powder and aliquots are stored at -20 °C
12. Domoic acid (Tocris Bioscience, catalog number: 0269); powder and aliquots are stored at -20 °C
13. NASPM trihydrochloride (Tocris Bioscience, catalog number: 2766); powder and aliquots are stored at -20 °C
14. ATPA [(RS)-2-Amino-3-(3-hydroxy-5-tert-butylisoxazol-4-yl)propanoic acid] (Tocris Bioscience, catalog number: 1107); powder and aliquots are stored at -20 °C
15. D-AP5 (D-2-Amino-5-phosphopentanoic acid) (Alomone Labs, catalog number: D-145); powder and aliquots are stored at -20 °C
16. HEPES [4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid] (AppliChem Panreac, catalog number: A3268); room temperature storage
17. Sodium chloride (NaCl) (USP, BP, Ph. Eur., JP) pure, pharma grade (AppliChem Panreac, catalog number: 141659); room temperature storage
18. Sodium phosphate dibasic (Na2HPO4) (Sigma-Aldrich, catalog number: S5136); room temperature storage
19. Potassium phosphate monobasic (KH2PO4) (Sigma-Aldrich, catalog number: P5156); room temperature storage
20. Potassium chloride (KCl) (USP, BP, Ph. Eur.) pharma grade (AppliChem Panreac, catalog number: 191494); room temperature storage
21. Sodium bicarbonate (NaHCO3) (Sigma-Aldrich, catalog number: P5761); room temperature storage
22. D-(+)-Glucose (Sigma-Aldrich, catalog number: G7021); room temperature storage
23. EDTA [2,2',2'',2'''-(Ethane-1,2-diyldinitrilo)tetraacetic acid] (AppliChem Panreac, catalog number: A5097); room temperature storage
24. Magnesium sulfate 7-hydrate (USP, BP, Ph. Eur.) pure, pharma grade (MgSO4·7H2O) (AppliChem Panreac, catalog number: 141404); room temperature storage
25. Calcium chloride (CaCl2) 1 M in H2O (Sigma-Aldrich, catalog number: 21115); store at 4 °C
26. Isoflurane (IsoNic) (Vetoquinol, 1,000 mg/g); store at 4 °C (the cap of the opened vial should be sealed with Parafilm M or another appropriate material)
27. Dimethyl sulfoxide (DMSO) (USP, BP, Ph. Eur.) (AppliChem Panreac, catalog number: 191954), store at room temperature in the dark
28. Trypan blue (Sigma-Aldrich, catalog number: 302643)
Solutions
1. Polyethyleneimine (PEI) solution (see Recipes)
2. Versene solution (see Recipes)
3. Hank’s balanced salt solution (HBSS) (see Recipes)
4. Neuron-glial cell culture growth medium (see Recipes)
5. Trypan blue 0.4% (w/v) solution (see Recipes)
Recipes
1. Polyethyleneimine (PEI) solution 1 mg/mL
Stock polyethyleneimine solution (Mn ~60,000; Mw 750,000) is jelly-like. Weigh the stock solution in a glass beaker, applying the substance on the beaker walls using a spatula. After weighing, add the required volume of double-distilled water and stir the solution for at least 30 min at room temperature using a magnetic stirrer. Then, filter the obtained PEI solution through a 0.22 μm membrane syringe filter for sterilization. Store the working solution at 4 °C and avoid freezing.
2. Versene solution
Reagent Final concentration
NaCl 137 mM
KCl 2.7 mM
KH2PO4 2 mM
Na2HPO4
EDTA
8 mM
0.6 mM
After EDTA addition, the solution should be heated to 40–50 °C at constant stirring to dissolve this chelator. To obtain a sterile Versen solution, it should be filtered through a 0.22 μm sterile syringe filter after dilution of all components. Double-distilled water is used to prepare the Versene solution. The prepared solution can be stored at room temperature or 4 °C.
3. Hank’s balanced salt solution
Reagent Final concentration
NaCl 137 mM
KCl 3 mM
Na2HPO4 0.35 mM
KH2PO4 1.25 mM
NaHCO3 4.2 mM
D-(+)-Glucose 10 mM
HEPES 10 mM
MgSO4·7H2O 0.8 mM
CaCl2 1.4 mM
Adjust pH to 7.35 using 10 M NaOH (the temperature of the solution during the pH adjustment must correspond to the temperature at which imaging experiments will be performed; in our studies, we perform all experiments at 28 °C). Since the neurons are sensitive to the pH of the extracellular solution, check the pH of HBSS before the experiments if HBSS is stored at 4 °C or more than one day at room temperature (slight alkalization is observed).
4. Neuron-glial cell culture growth medium
Reagent Final concentration
Neurobasal A-medium n/a
50× B27 supplement 2% (v/v)
L-glutamine 0.5 mM
100× Penicillin-streptomycin 1% (v/v)
L-glutamine is unstable in aqueous solutions. Add L-glutamine immediately before the cell culture preparation or use stable analogs. Due to the instability of some components, we recommended preparing small portions of neuron-glial cell culture growth medium (50–100 mL) and storing them at 4 °C for no more than 1 week.
5. Trypan blue 0.4% (w/v) solution
Trypan blue 0.4% working solution is prepared by dissolving the weighed powder in phosphate-buffered saline (PBS). PBS composition is listed below (pH 7.35). Before the application, the solution was filtered through a syringe filter with a pore size of 0.22 μm to remove undissolved aggregates.
Reagent Final concentration
NaCl 137 mM
KCl 2.7 mM
KH2PO4 2 mM
Na2HPO4 8 mM
Laboratory supplies
1. Cell culture dishes 35 mm (SPL Lifesciences, catalog number: 11035)
2. Cell culture dishes 60 mm (SPL Lifesciences, catalog number: 11060)
3. Cell culture dishes 150 mm (SPL Lifesciences, catalog number: 10151)
4. Sterile microcentrifuge tube 1.5 mL (GenFollower, catalog number MCTB015)
5. Sterile centrifuge tubes 50 mL (GenFollower, catalog number B-2SCTB50YE)
6. Low-retention pipette tips 200 μL (GenFollower, catalog number E-FTR200-L-S)
7. Low-retention pipette tips 1000 μL (GenFollower, catalog number E-FTR1000-L-S)
8. Round cover glasses 25 mm (VWR, catalog number 630-2122)
9. Syringe filter 0.22 μm (Membrane Solutions, catalog number MS-SFPES025022SI)
10. Cover glasses 24 × 24 (Minimed, catalog number 12003316)
Equipment
1. Thermoshaker (Biosan, model: TS-100, catalog number BS-010120-AAI) with thermoblock SC-18/02 (Biosan, catalog number BS-010120-CK)
2. High-speed centrifuge (DLab, model: D2012, catalog number 9032002121)
3. Laminar flow hood (Lamsystems, model: BMB-II-"Laminar-S."-1,2 NEOTERIC, catalog number 2E-B.001-12)
4. Trinocular inverted microscope (Nikon, model: Eclipse TS100)
5. Stereo microscope (Leica Microsystems, model: EZ4D)
6. CO2 incubator (N-Biotek, model: NB-203XL)
7. Epifluorescent inverted microscope (Leica Microsystems, DMI 6000B) with CCD-camera (Hammamatsu, model: 9100C) and illuminator (Leica Microsystems, model: EL6000)
8. Analytical scale (Ohaus, model: Explorer EX324)
9. pH meter (Mettler Tolledo, model: S20 SevenEasy)
10. Magnetic stirrer with a hot plate (Biosan, model: MSH-300)
11. Bidistiller (GFL, model: 2104)
12. Multi-output animal anesthesia machine (RWD Life Science, model: R550)
13. Drying and heating chamber with natural convection (Binder Inc., model: ED23)
14. Neubauer chamber (JVLAB, catalog number: 1103)
15. Instruments for tissue extraction and preparation: dental curved tweezers 150 mm (catalog number: 10-94), dental spatula (catalog number: 10-97), and medical blunt-pointed straight scissors (catalog number: 4-26) from Mozhaisk Medical Supplies Factory; curved scissors (catalog number: ТН-06-041-11,3) and scalpel (catalog number: TC-02-051-15) from Tumbotino Medical Supplies Factory; sharp-pointed straight (catalog number: ST-14) and curved (catalog number: ST-15) tweezers from NAGARAKU, Shanghai Xijian Electronic Technology Co., Ltd.
Software and datasets
1. ImageJ v1.53s (https://imagej.net/ij/, May 2022)
2. OriginLab Pro 2016 version b9.3.226 (commercially available software, license required; other free software for plot creation can be used as an alternative, October 2015)
Procedure
A. Hippocampal cell culture preparation
1. Coat the round cover glasses with polyethyleneimine solution.
a. Sterilize the glasses in a dry heat sterilizer (or using a laboratory burner).
Note: Sterilization in a dry heat sterilizer should be performed for 2 h at a temperature of 150 °C. In the case of the laboratory burner, use tweezers and hold a cover glass in the flame for no more than 1 s each side. After that, hold for 10–20 s away from the flame to allow it to cool. If the cover glass is overheated, it can crack during cooling. Hence, sterilization with a dry heat sterilizer is preferable.
b. Coat sterile glasses with 1–2 mL of polyethyleneimine solution and leave it in a laminar hood for 1 h (fan should be turned off to prevent drying; on-board UV lamp can be turned on).
Note: You can simultaneously coat dozens of cover glasses placed in large sterile Petri dishes (≥120–150 mm in diameter) or another appropriate bath. For these purposes, both dishes and lids can be used. Approximately 15–18 cover glasses (25 mm diameter) can be arranged in a 150 mm Petri dish.
c. Wash the glasses three times with sterile (boiled for 15 min and cooled to room temperature) double-distilled water and leave them for 12 h in a laminar hood to dry completely.
Note: If the Petri dishes are used for the arrangement of the cover glasses during PEI treatment, sterile water can be added to the dish during the washing procedures. The volume of water used for each wash step depends on the used bath but, to efficiently wash the cover glasses, the level of water in the bath has to be 3–5 mm higher than the surface of the cover glasses.
d. Place the PEI-treated dry glasses in the sterile 35 mm cell culture dishes.
2. Before the hippocampus extraction, prepare the sterile instruments (medical scissors, scalpel, dental spatula, and tweezers) and a small bath with ice.
3. Place in the bath the 60 mm cell culture dish with the Versene solution (8–10 mL) and two microcentrifuge tubes (one with 1 mL of neuron-glial cell culture growth medium and the second one with Versene solution).
4. To euthanize the animal, put it in the preconditioning (induction) chamber and fill the chamber with isoflurane (5%) or another appropriate inhaled anesthetic.
Note: The incubation time depends on the anesthesia used and the weight and strain of the used animal.
5. After incubation, take the euthanized animal from the chamber and thoroughly treat it with 70% ethanol (it is better to use a pulverizer for this purpose).
6. Then, using sharp medical scissors, quickly decapitate the animal.
7. Carefully cut the skull using sharp scissors (curved nail scissors are recommended for this procedure).
8. Extract the brain using a dental spatula and put it into the 60 mm cell culture dish with the ice-cold Versene solution.
9. Cut hemispheres along the line from the cerebellum to the olfactory bulb with a scalpel.
Note: It is better to use curve tweezers to hold the brain during making the cut and further procedures.
10. Remove the parts of the hippocampus from both hemispheres and put them into the 1.5 mL microcentrifuge tube with ice-cold Versene solution.
Note: We recommend using a dental spatula and tweezers to remove the hippocampus.
11. Cut the tissue in the tube into 0.5–2 mm fragments using sterile scissors.
12. Replace the Versene solution in the tube with 500 μL of 1% trypsin solution.
Note: To obtain the working trypsin solution, dilute the stock in Versene.
13. Incubate the hippocampal fragments with trypsin at 37 °C at constant stirring for 10 min (stirring rate of 500 rpm, 0.6× g).
Note: A laboratory shaker with a heater can be used for this purpose.
14. Remove the trypsin solution and wash the tissue fragments two times with 500 μL of cold neuron-glial cell culture growth medium to inactivate trypsin.
Note: Avoid the trituration of tissue fragments when replacing the medium in the tube. Pipette tips produced from plastic with low retention are recommended since the trypsinized tissue fragments are sticky.
15. Add 1 mL of room-temperature neuron-glial cell culture growth medium and gently triturate tissue fragments using a 1 mL pipette tip.
16. Wait for 30–60 s until the non-triturated fragments sediment; then, carefully remove them.
17. Carefully remove the non-sedimented film-like tissue fragments using a 200 μL pipette tip.
18. Centrifuge the obtained cell suspension at 2,000× g for 3 min at room temperature.
19. Remove the supernatant and resuspend the pellet to obtain a cell suspension with a cell quantity of 106107 cells/mL.
Note: The cells can be counted manually with different counting chambers or automatically using cell counters. We used the Neubauer chamber for cell counting. For the staining, the “stock” cell suspension was diluted 10 times with neurobasal medium to expedite the counting procedure, and this dilution factor was considered in calculations of average cell quantity. Before counting, the diluted cell suspension was mixed in proportion 1:1 with 0.4% trypan blue solution (5 min staining).
20. Dilute the obtained suspension if necessary and drop the aliquots of the cell suspension on the PEI-treated round cover glasses. The optimal cell density is 30,000–50,000 cells/mm2.
Note: To concentrate the cells in the restricted area of the cover glass, we use the sterile glass cylinders placed on the cover glasses lying in 35 mm cell culture dishes (Figure 1A). The height used in the study cylinders was 6 mm, and the internal diameter was 5 mm; the working volume ≈ 100 μL. The cylinders can be produced from glass serological pipettes, for instance. After cell attachment, the cylinders are carefully removed with the curved tweezers. Twenty-four hours after the preparation, the representative image of the hippocampal cell culture (Figure 1B) demonstrates the average cell density that can be achieved using glass cylinders.
Figure 1. Hippocampal cell attachment. A. Glass cylinders used during cell culture preparation. B. Brightfield images of a representative cell culture 24 h after cell attachment.
21. Put the cell culture dishes with the cylinders in the CO2 incubator for 30–60 min for cell attachment.
22. Remove the dishes with the cell cultures and add 2 mL of the neuron-glial cell culture growth medium (temperature ≈ 37 °C) in each cell culture dish. If the glass cylinders were used, carefully remove them (use curve tweezers) before the addition of the growth medium.
Note: The average passage of the hippocampal cell cultures obtained from one animal includes 12–14 cell cultures (cell culture dishes with the PEI-treated glasses covered with the hippocampal cells).
23. Cultivate the cultures in a CO2 incubator at 37 °C in an atmosphere containing 5% CO2 (humidity ≥ 95%) for 12–14 days and use in experiments.
Note: Frequent cell culture medium replacement affects the quality of neuronal networks in the cell culture. Replace 1/2 of the medium volume at 5–6 DIV (days in vitro) and keep the humidity in the incubator at ≥ 95% (this is important).
B. Staining of the cultures with Fura-2
1. Dissolve Fura-2 AM stock solution (1 mM in DMSO) to a working concentration of 3 μM and incubate the cell cultures with the working solution for 40 min at 28 °C in the dark.
Note: To decrease the consumption of Fura-2, the glasses with the cell cultures can be transferred from the cell culture dishes to their lids and covered with 200 µL of the working solution. This volume is enough for cell staining.
2. Remove the dishes with 12–14 DIV neuron-glial cell cultures from the CO2 incubator, wash them twice with HBSS solution (2 mL per wash), wait for 10 min, and wash once again (2 mL).
Note: The pause between washes is required to allow the non-esterified probe to flow from the cells to the extracellular medium.
3. Mount the cover glass with the cell culture into the chamber for measurements.
4. Choose the area with the monolayer to record the signal from the maximal number of cells. The optimal ratio between magnification and the number of cells in the view field can be achieved with 10.0× and 20.0× objectives (Figure 2A).
5. Use a perfusion system providing a 10 mL/min flow rate to effectively and quickly replace the medium in the microscope chamber.
Figure 2. Fura-2 imaging. A. Representative image of Fura-2-stained rat hippocampal cell culture. Objective HC PL FLUOTAR 10.0 × 0.30 DRY with an additional 1.6 magnification lens. Red arrows show the neurons whose changes in intracellular Ca2+ concentration are shown in panel B. Images show the fluorescence of cells before (20 s) and during (60 s) the KCl application (see panel B). Excitation 340 nm: maximum of the excitation spectrum of Ca2+-bound form of Fura-2; 387 nm: maximum of the excitation spectrum of Ca2+-free form of Fura-2. B. Changes of intracellular Ca2+ concentration in cells of the neuron-glial culture in response to KCl (35 mM) application. Red curves correspond to neurons, while green curves correspond to glial cells.
C. Identification of neurons in hippocampal cell culture
1. To visualize the neurons stained with a Ca2+-sensitive fluorescent probe (Fura-2 AM in this case), perform the application of 35 mM KCl.
Note: As shown in previous works, KCl application (35 mM) depolarizes the neurons and stimulates the release of neurotransmitters, particularly glutamate. Depolarization induced by KCl and activation of ionotropic glutamate receptors (AMPARs, NMDARs, and KARs) promotes the opening of voltage-gated Ca2+ channels (VGCC) [13,14], mediating Ca2+ inflow into the soma of neurons. These Ca2+ changes are detected with different Ca2+-sensitive probes, such as Fura-2, Fluo-3, Fluo-4, or Fluo-8. Neurons quickly restore calcium homeostasis after KCl application. Therefore, this method of identification of all neurons in a view field can be used in routine calcium imaging experiments.
2. The cells responding with a sustained increase in intracellular Ca2+ concentration ([Ca2+]i) to KCl application are neurons (Figure 2B).
Note: Using a combination of live cell fluorescent calcium imaging and post-vital immunostaining, we have previously demonstrated that cells responding to KCl application contain the neuronal marker, NeuN [15].
D. Identification of neurons containing Ca2+-permeable AMPA receptors
1. To identify neurons containing the calcium-permeable AMPA receptors, perform the application of selective AMPAR agonist (5-fluorowillardiine, AMPA, etc.) in the presence of non-selective blocker of VGCC (verapamil or diltiazem, for example), antagonists of NMDA and kainate receptors [16].
Note: As shown in Figure 3A, [Ca2+]i elevation in neurons in the presence of AMPAR agonists is mediated in most neurons by VGCC, NMDARs, and KARs. The sustained elevation in the presence of the appropriate antagonists/blockers remains only in a minor population of neurons.
2. The cells responding with sustained [Ca2+]i elevation to the application of AMPAR agonists in the presence of VGCC blockers and NMDAR and KAR antagonists can be attributed to neurons containing calcium-permeable AMPA receptors (CP-AMPAR-neurons).
Note: This [Ca2+]i elevation is sensitive to the antagonist of Ca2+-permeable AMPARs, NASPM (Figure 3B).
Figure 3. Identification of neurons containing CP-AMPARs. A. Responses of neurons in rat hippocampal neuron-glial cell cultures to the application of selective AMPAR agonist, 5-fluorowillardiine (FW, 500 nM) in the absence and presence of the blocker of voltage-gated calcium channels (verapamil, 300 μM) and NMDAR and KAR antagonists (D-AP5, 10 μM and UBP310, 10 μM, respectively). Red curves: neurons containing CP-AMPARs; grey curves: other neurons. B. Effect of NASPM (selective antagonist of CP-AMPARs, 30 μM) on the [Ca2+]i elevation induced by AMPAR activation in neurons containing CP-AMPARs.
D. Identification of neurons containing Ca2+-permeable KA receptors
1. To identify neurons containing the calcium-permeable KA receptors, perform two applications of KAR/AMPAR agonist, domoic acid (DoA, 200–500 nM), in the presence and absence of NASPM (30 μM).
Note: We have shown in our previous works that in rat hippocampal neuron-glial cultures, the neurons containing CP-AMPARs and CP-KARs are GABAergic [16,17], and their selective activation suppresses the activity of the innervated glutamatergic neurons. Due to insufficient GABA-mediated innervation, GABAergic neurons [16], including neurons containing CP-KARs, respond earlier to depolarizing stimuli. To eliminate the effects associated with GABA-mediated inhibition and the identification of neurons containing CP-KARs, DoA applications should be performed in the presence of GABA(A)R antagonists [the contribution of GABA(B)R is less pronounced in this case]. We used bicuculline (10 μM) for this purpose, but other GABA(A)R antagonists, such as gabazine or picrotoxin, can also be used. Notably, the application of GABA(A)R antagonists induces epileptiform activity manifested as oscillations of [Ca2+]i.
2. The neurons insensitive to NASPM (Figure 4, green curves) can be attributed to neurons containing calcium-permeable KARs.
Note: We have demonstrated that neurons containing CP-KARs and neurons containing CP-AMPARs also respond with [Ca2+]i elevation to the application of selective agonist of GluK1/GluK3 containing KARs, ATPA (300–500 nM) [16]. Since ATPA demonstrates lower excitotoxicity than DoA or selective AMPAR agonists, ATPA-induced [Ca2+]i elevations can be considered an additional marker of GABAergic neurons containing CP-KARs or CP-AMPARs.
Figure 4. Identification of neurons containing CP-KARs. A. Responses of representative neurons to AMPAR/KAR antagonist, domoic acid (DoA 200 nM) in the presence or absence of NASPM (30 μM). Both DoA applications were performed in the presence of GABA(A)R antagonist, bicuculline (10 μM). Green curves correspond to neurons insensitive to NASPM and expressing CP-KARs. The neurons in which the amplitude of the sustained DoA-induced [Ca2+]i elevation decreases in the presence of NASPM (red curves) contain CP-AMPARs. Grey curves: other neurons without CP-AMPARs. B. Averaged responses of neurons from panel A.
Data analysis
1. The series of fluorescent images can be analyzed with free software ImageJ.
2. To subtract background noise, use the Subtract Background function in ImageJ (Process → Subtract Background). The variable parameter Rolling ball radius should be chosen in the range of 30–50 pixels (depending on magnification).
3. To calculate time-lapse changes in the fluorescence intensity of cells, place ROIs on the soma without covering processes to avoid capturing the background fluorescence (Figure 5).
Figure 5. Analysis of representative Fura-2 images. A, B. Fura-2 fluorescence upon 387 nm excitation (Ca2+-free form of the probe) before (A) and during (B) KCl application. C. 340/387 Fura-2 ratio during KCl application; “Fire” lookup table. Yellow circles demonstrate the regions of interest (ROIs) where the average signal intensity was measured. Scale bar 20 μm.
4. In the case of ratiometric probes, such as used Fura-2, the series of images corresponding to the channels with excitation 340 nm have to be divided by the appropriate series of images corresponding to the channel with excitation 387 nm (Process → Image Calculator; 32-bit option should be chosen).
5. To calculate the mean Fura-2 340/387 ratio (reflects the changes in [Ca2+]i) for cells in a view field, perform the command in ROI Manager More → Multi Measure. Set Mean Grey Value as the calculated parameter (Analyze → Set Measurements).
6. Copy or export obtained results and draw plots using any appropriate software (e.g., OriginLab Pro).
7. In the case of non-ratiometric Ca2+-sensitive fluorescent probes (Fluo-4 family or others), use similar algorithms excluding step 4.
Validation of protocol
This protocol or parts of it have been used and validated in the following research articles:
Zinchenko et al. [17]. Properties of GABAergic Neurons Containing Calcium-Permeable Kainate and AMPA-Receptors. Life (Basel) (Figures 1–5).
Gaidin et al. [16]. A novel approach for vital visualization and studying of neurons containing Ca2+ -permeable AMPA receptors. J Neurochem (Figures 1–6).
Maiorov et al. [10]. Peculiarities of ion homeostasis in neurons containing calcium-permeable AMPA receptors. Arch Biochem Biophys (Figures 1–4).
Zinchenko et al. [18]. Participation of calcium-permeable AMPA receptors in the regulation of epileptiform activity of hippocampal neurons. Front synaptic neurosci (Figure 3).
General notes and troubleshooting
General notes
1. The proposed methods can be used not only in cell cultures but also in brain slices.
2. Any Ca2+-sensitive fluorescent cell-permeant probes can be used in the experiments.
3. The method allows combining with electrophysiological measurements or immunostaining.
4. The described approaches allow the expressed identification of many neurons in a view field.
5. Since permeability for Ca2+ is determined in the case of AMPARs and KARs not only by the subunit composition but also by Q/R editing of pre-mRNA encoding GluA2, GluK1, GluK2 subunits, we suppose that the described approach allows identification of receptors consisting of non-edited subunits.
6. We suppose that the suggested approaches allow the identification only of neurons containing a significant number of CP-AMPARs and CP-KARs. If the percentage of the calcium-permeable receptors in a neuron is low, the signals of the Ca2+-sensitive probes can be negligible.
Troubleshooting
Problem 1: Low-amplitude response to DoA (or the absence of the response) or slow restoration of [Ca2+]i after washout.
Possible cause: Inappropriate concentration of the agonist.
Solution: Change the concentration (the optimal ranges are listed above).
Problem 2: Absence of response to AMPAR agonists in the presence of NMDAR, KAR antagonists, and VGCC blockers.
Possible cause: Disturbances of the neuronal network formation and death of neurons at early stages of the cultivation.
Solution: Increase cell density during the culture preparation to improve the viability of neurons and promote synaptogenesis and maturation. Hold the humidity in a CO2 incubator at ≥ 95% to prevent excessive evaporation from the cell culture dishes and shift the osmolarity of the neuron-glial culture medium.
Acknowledgments
The previously obtained results underlying the proposed methods of identification were supported by the Ministry of Science and Higher Education of the Russian Federation in the framework of state assignment of PSCBR RAS 075-01512-22-02/075-00609-24-01 (No 1022080100047-5-1.6.4, Neuroprotective drugs of a new generation) (animal facility resources, technical support for the experimental studies and cell culture preparations) and by the Science Committee of the Ministry of Science and Higher Education of the Republic of Kazakhstan (Grant No. AP19678607; Grant no. AP05133528; Grant no. AP19680470). The current work was funded by a grant from the Science Committee of the Ministry of Science and Higher Education of the Republic of Kazakhstan Grant No. AP19678607.
Competing interests
The authors declare no conflict of interest.
Ethical considerations
All animal procedures were approved by the Bioethics Committee of the Institute of Cell Biophysics (ICB) and carried out according to Act708n (August 23, 2010) of the Russian Federation National Ministry of Public Health, which states the rules of laboratory practice for the care and use of laboratory animals, and the Council Directive 2010/63 EU of the European Parliament on the protection of animals used for scientific purposes. ICB RAS Animal Facility provided the animals for experiments in accordance with the applications approved by the Commission on Biosafety and Bioethics of Institute of Cell Biophysics (Permission No. 6, 12 December 2017; Permission No. 2, 12 June 2020, Permission No. 3, 12 April 2021; Permission No. 4, 17 July 2021, Permission No. 3, February 12, 2022; Permission No. 4, 17 June 2022, Permission No. 3, March 12, 2023).
References
Park, P., Kang, H., Sanderson, T. M., Bortolotto, Z. A., Georgiou, J., Zhuo, M., Kaang, B. K. and Collingridge, G. L. (2018). The Role of Calcium-Permeable AMPARs in Long-Term Potentiation at Principal Neurons in the Rodent Hippocampus. Front Synaptic Neurosci. 10: e00042. https://doi.org/10.3389/fnsyn.2018.00042
Sanderson, J. L., Gorski, J. A. and Dell’Acqua, M. L. (2016). NMDA Receptor-Dependent LTD Requires Transient Synaptic Incorporation of Ca 2+ -Permeable AMPARs Mediated by AKAP150-Anchored PKA and Calcineurin. Neuron. 89(5): 1000–1015. https://doi.org/10.1016/j.neuron.2016.01.043
Gaidin, S. G. and Kosenkov, A. M. (2023). Calcium-permeable AMPA receptors. Neural Regen Res. 18(12): 2669–2670. https://doi.org/10.4103/1673-5374.373714
Guo, C. and Ma, Y. Y. (2021). Calcium Permeable-AMPA Receptors and Excitotoxicity in Neurological Disorders. Front Neural Circuits. 15: e711564. https://doi.org/10.3389/fncir.2021.711564
Azarnia Tehran, D., Kochlamazashvili, G., Pampaloni, N. P., Sposini, S., Shergill, J. K., Lehmann, M., Pashkova, N., Schmidt, C., Löwe, D., Napieczynska, H., et al. (2022). Selective endocytosis of Ca 2+ -permeable AMPARs by the Alzheimer’s disease risk factor CALM bidirectionally controls synaptic plasticity. Sci Adv. 8(21): eabl5032. https://doi.org/10.1126/sciadv.abl5032
Martinez, T. P., Larsen, M. E., Sullivan, E., Woolfrey, K. M. and Dell’Acqua, M. L. (2024). Amyloid-β-Induced Dendritic Spine Elimination Requires Ca2+-Permeable AMPA Receptors, AKAP-Calcineurin-NFAT Signaling, and the NFAT Target Gene Mdm2. eNeuro. 11(3). https://doi.org/10.1523/eneuro.0175-23.2024
Kobylecki, C., Cenci, M. A., Crossman, A. R. and Ravenscroft, P. (2010). Calcium‐permeable AMPA receptors are involved in the induction and expression of l‐DOPA‐induced dyskinesia in Parkinson’s disease. J Neurochem. 114(2): 499–511. https://doi.org/10.1111/j.1471-4159.2010.06776.x
Zhang, J., Wang, Y., Sun, Y. N., Li, L. B., Zhang, L., Guo, Y., Wang, T., Yao, L., Chen, L., Liu, J., et al. (2019). Blockade of calcium-permeable AMPA receptors in the lateral habenula produces increased antidepressant-like effects in unilateral 6-hydroxydopamine-lesioned rats compared to sham-lesioned rats. Neuropharmacology. 157: 107687. https://doi.org/10.1016/j.neuropharm.2019.107687
Coombs, I., Bats, C., Sexton, C. A., Studniarczyk, D., Cull-Candy, S. G. and Farrant, M. (2023). Enhanced functional detection of synaptic calcium-permeable AMPA receptors using intracellular NASPM. eLife. 12: e66765. https://doi.org/10.7554/elife.66765
Maiorov, S., Zinchenko, V., Gaidin, S. and Kosenkov, A. (2021). Potential mechanism of GABA secretion in response to the activation of GluK1-containing kainate receptors. Neurosci Res. 171: 27–33. https://doi.org/10.1016/j.neures.2021.03.009
Lalanne, T., Oyrer, J., Mancino, A., Gregor, E., Chung, A., Huynh, L., Burwell, S., Maheux, J., Farrant, M., Sjöström, P. J., et al. (2015). Synapse‐specific expression of calcium‐permeable AMPA receptors in neocortical layer 5. J Physiol. 594(4): 837–861. https://doi.org/10.1113/jp271394
Rajasekaran, K., Todorovic, M. and Kapur, J. (2012). Calcium‐permeable AMPA receptors are expressed in a rodent model of status epilepticus. Ann Neurol. 72(1): 91–102. https://doi.org/10.1002/ana.23570
Laryushkin, D. P., Maiorov, S. A., Zinchenko, V. P., Gaidin, S. G. and Kosenkov, A. M. (2021). Role of L-Type Voltage-Gated Calcium Channels in Epileptiform Activity of Neurons. Int J Mol Sci. 22(19): 10342. https://doi.org/10.3390/ijms221910342
Gaidin, S. G., Zinchenko, V. P., Teplov, I. Y., Tuleukhanov, S. T. and Kosenkov, A. M. (2019). Epileptiform activity promotes decreasing of Ca2+ conductivity of NMDARs, AMPARs, KARs, and voltage-gated calcium channels in Mg2+-free model. Epilepsy Res. 158: 106224. https://doi.org/10.1016/j.eplepsyres.2019.106224
Gaidin, S. G., Zinchenko, V. P., Sergeev, A. I., Teplov, I. Y., Mal'tseva, V. N. and Kosenkov, A. M. (2019). Activation of alpha‐2 adrenergic receptors stimulates GABA release by astrocytes. Glia. 68(6): 1114–1130. https://doi.org/10.1002/glia.23763
Gaidin, S. G., Maiorov, S. A., Laryushkin, D. P., Zinchenko, V. P. and Kosenkov, A. M. (2022). A novel approach for vital visualization and studying of neurons containing Ca2+‐permeable AMPA receptors. J Neurochem. 164(5): 583–597. https://doi.org/10.1111/jnc.15729
Zinchenko, V. P., Kosenkov, A. M., Gaidin, S. G., Sergeev, A. I., Dolgacheva, L. P. and Tuleukhanov, S. T. (2021). Properties of GABAergic Neurons Containing Calcium-Permeable Kainate and AMPA-Receptors. Life 11(12): 1309. https://doi.org/10.3390/life11121309
Zinchenko, V. P., Teplov, I. Y., Kosenkov, A. M., Gaidin, S. G., Kairat, B. K. and Tuleukhanov, S. T. (2024). Participation of calcium-permeable AMPA receptors in the regulation of epileptiform activity of hippocampal neurons. Front Synaptic Neurosci. 16: e1349984. https://doi.org/10.3389/fnsyn.2024.1349984
Article Information
Publication history
Received: Oct 7, 2024
Accepted: Dec 10, 2024
Available online: Jan 9, 2025
Published: Feb 5, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
How to cite
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Neuroscience > Neuroanatomy and circuitry > Fluorescence imaging
Neuroscience > Cellular mechanisms > Synaptic physiology
Cell Biology > Cell imaging > Fluorescence
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52 | https://bio-protocol.org/en/bpdetail?id=52&type=1 | # Bio-Protocol Content
Improve Research Reproducibility
A Bio-protocol resource
This is an In Press version of the protocol that has not yet been assigned to an issue.
Peer-reviewed
Standard DNA Cloning
Fanglian He
In Press
Published: Apr 5, 2011
DOI: 10.21769/BioProtoc.52 Views: 65502
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Abstract
This protocol describes general cloning steps from preparation of both vector and insert DNA to the ligation reaction.
Materials and Reagents
Luria-Bertani broth (LB) medium: Bacto-tryptone (BD Biosciences), yeast extract (BD Biosciences)
Antibiotics (Sigma-Aldrich/Thermo Fisher Scientific)
QIAGEN Plasmid Purification Handbook (QIAGEN)
SeaKem® LE Agarose (Cambrex)
Plasmid Prep Kit (QIAGEN /Fermentas)
PCR Clean-up kit (QIAGEN /Fermentas)
Restriction enzymes (New England Biolabs)
Alkaline Phosphatase: Calf intestinal alkaline phosphatase (CIP) (New England Biolabs, catalog number: M0290) or Shrimp Alkaline Phosphatase (SAP) (Promega Corporation, catalog number: M8201 )
Ligase enzyme (New England Biolabs)
DNA ladder
NaCl
LB broth media (see Recipes)
Ligation reaction (see Recipes)
Equipment
Nanodrop (Thermo Scientific)
Procedure
Preparing vector DNA for cloning:
Depending on the copy number of the vector plasmid, decide if you need the Mini-prep, Midi-prep, or Maxi-prep kit. If it is a high copy (>10 copies/cell) plasmid, plasmid DNA can be prepared by using the Mini-prep kit. If it is a low copy (<10 copies/cell) plasmid, use the Midi-prep or Maxi-prep kit.
Grow E. coli cell culture carrying vector plasmid in LB liquid medium with appropriate antibiotics at 37 °C overnight.
Follow QIAGEN Plasmid Purification Handbook to obtain DNA. If plasmid DNA does not need to be purified, and to be more economical, plasmid DNA can be extracted without using a plasmid prep kit (See protocol “Plasmid DNA extraction from E. coli using alkaline lysis method”).
Estimate plasmid DNA concentration using one of the following two ways:
Load 2-3 μl plamid DNA and a DNA ladder on a DNA agarose gel and estimate DNA according to the DNA marker.
Easier and more accurate way is to measure DNA using Nanodrop if it is available.
Digest 2-5 μg vector DNA using restriction enzymes needed for the insert DNA. To make sure the vector is completely digested, extra enzyme and long incubation may be needed.
To reduce the chance of self-ligation, dephosphorylate the 5′ phosphorylated ends of the digested vector with alkaline phosphatase.
Note: If the shrimp alkaline phosphatase (SAP) is used, then add 2 μl SAP directly to 100 μl digest solution, incubate at 37 °C for 1 h, then inactivate SAP at 65 °C for 10 min. If the calf intestinal alkaline phosphatase (CIP) is used, then add 5 μl CIP enzyme to 100 μl digestion solution, incubate at 37 °C for 1 h, then inactivate SAP at 65 °C for 30 min.
Perform gel purification of digested vector DNA.
Preparing insert DNA for cloning:
Obtain insert DNA from digestion of plasmid DNA.
Extract plasmid DNA as described above.
Digest plasmid DNA with appropriate restriction enzymes.
Perform gel purification of insert DNA.
Generate insert DNA from PCR product.
Design primers using a free a good quality program online (e.g., http://frodo.wi.mit.edu/primer3/) containing desired cloning sites with several of bases flanking their recognition sequences (http://www.neb.com/nebecomm/tech_reference/restriction_enzymes/ cleavage_olignucleotides.asp).
Amplify insert DNA from a template by PCR, and clean up PCR product by PCR clean-up kit.
Digest PCR product with the corresponding restriction enzymes. Or, first clone PCR product to pGEM T-easy vector, and then generate insert DNA from the resulting plasmid.
Perform gel purification of insert DNA.
Estimate DNA concentration.
Ligation of insert and vector:
Usually (particularly for blunt end ligation), need more insert DNA than vector: 1 mole of vector normally needs 5 or more moles of insert (see protocol “DNA molecular weight calculation”).
Control ligation: To determine background clones arising from self-ligation of inefficiently phosphatased vector, set a parallel ligation in the absence of insert DNA.
Transform 1 μl ligation reaction to competent cell by electroporation or chemical method.
Colony PCR to screen for plasmids carrying the correct inserts and then confirm the result by digestion and sequencing of the plasmid.
Recipes
1 liter of LB broth media
10 g Bacto-tryptone
5 g yeast extract
10 g NaCl
Add ddH2O to get volume 1 L
Sterilize by autoclaving.
Ligation reaction
X μl DNA vector( ~20 ng)
Y μl insert (~100-1,000 ng)
2 μl 10x buffer
1 μl T4 DNA ligase
To 20 μl H2O
--------
20 μl total
Article Information
Copyright
© 2011 The Authors; exclusive licensee Bio-protocol LLC.
How to cite
Category
Molecular Biology > DNA > DNA cloning
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5,200 | https://bio-protocol.org/en/bpdetail?id=5200&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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Peer-reviewed
An Effective and Safe Maize Seed Chipping Protocol Using Clipping Pliers With Applications in Small-Scale Genotyping and Marker-Assisted Breeding
BZ Brian Zebosi
JS John Ssengo
LG Lander F Geadelmann
EU Erica Unger-Wallace
EV Erik Vollbrecht
Published: Vol 15, Iss 3, Feb 5, 2025
DOI: 10.21769/BioProtoc.5200 Views: 65
Reviewed by: Samik Bhattacharya Anonymous reviewer(s)
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Abstract
In applications such as marker-assisted breeding and positional cloning, tissue sampling and plant tracking are vital steps in the genotyping pipeline. They enable the identification of desirable seedlings, saving time and reducing the cost, space, and handling required for growing adult plants, especially for greenhouses and winter nurseries. Small-scale marker-assisted selection laboratories rely heavily on leaf-based genotyping, which involves over-planting large, segregating populations followed by leaf sampling, genotyping, and backtracking to identify desired individuals, which is costly and laborious. Thus, there is a need to adopt seed-based genotyping to reduce costs and save time. Therefore, we developed a safe and cheap seed-chipping protocol using clipping pliers to chip seeds to genotype before planting. To identify a cost-effective and high-throughput DNA extraction method, we tested four extraction methods and assessed the quality of the seed DNA using PCR. For three of the methods, seed-based DNA was of comparable quality to DNA extracted from leaf punches. We also compared seed- and leaf-derived DNA from the same individuals in a segregating population to test for genotyping miscalls that could arise due to the presence of maternally derived pericarp in the seed samples. Out of 43 potential instances, we found zero miscalled samples and, therefore, no evidence supporting consequential pericarp inclusion. Germination rates of chipped and unchipped seeds were the same for the inbreds tested, B73 and Mo17. However, chipped seeds grew slower until ~14 days after sowing. Overall, seed sampling using clipping pliers provides a simple, reliable, and high-throughput method to identify specific genotypes before planting.
Key features
• Provides a quick, safe, and cheap sampling technique for maize kernels that may also be suitable for other plants with relatively large seeds.
• Includes procedures and materials to track and organize samples within and across batches involving tens to thousands of seeds.
• Seeds can be sampled and genotyped relatively quickly for planting; in one day, 384 seeds can be sampled, processed for DNA, and genotyped by PCR.
Keywords: Maize Genotyping DNA extraction Seed chipping Kernel Endosperm Grain
Background
Over the last decades, marker-assisted selection (MAS) has become integral to most plant breeding systems because quantitative trait loci (QTL) and molecular markers linked to desired traits have enhanced the selection efficiency of desirable plants and shortened breeding cycles [1]. Unlike commercial breeding companies with highly automated and streamlined MAS and trait-discovery programs, public-sector breeding programs and small-scale genotyping laboratories rely on laborious and time-consuming tissue sampling and DNA extraction procedures [2]. Compared to the proprietary seed-based MAS programs utilized by industry scientists, public-sector genotyping systems that rely heavily on leaf-based DNA genotyping may be particularly resource- and time-demanding [1] as the marker genotyping process must be coupled to the sowing and germinating of seeds, caring for seedlings, tracking of live plants to static samples, and the immediate interpretation of data in order to identify, retain, and rear the desired, growing plants for their intended purpose. Thus, to improve the efficiency of MAS and related approaches, there is a need to adopt seed-based genotyping as a separate, pre-planting process, which reduces overall labor costs and saves space and time through selection of desirable genotypes during the off-season [1,2]. During trait discovery, seed-based DNA genotyping ensures that seeds with desirable traits are selected and planted, thus saving time and costs associated with greenhouse use and winter fields. In addition to tissue sampling, optimizing rapid and cost-effective DNA extraction methods to screen large populations will be essential for a successful MAS and genotyping pipeline.
Seed-based DNA genotyping methods have been implemented in small-scale maize breeding programs and genotyping laboratories but are still ineffective based on safety and scalability [2,3]. One seed-based DNA genotyping method is based on soaking maize kernels in water before endosperm chipping [2]. The caveats of this procedure include susceptibility to fungal contamination and low seed germination if kernels are not dried and stored well. Another common method uses a razor blade for seed chipping [3], which presents injury hazards and is time-consuming, thus making it impractical to quickly chip a large population. In this study, we used clipping pliers, a safe alternative to previous chipping methods that is scalable to high-throughput applications. This seed-chipping method did not significantly impact germination rates for maize B73 and Mo17 seeds, although seedlings from chipped seeds had reduced vigor that was evident for 14 days after planting, after which they appeared comparable to their non-chipped sibling seedlings. In addition, we tested four DNA extraction methods, based on each of cetyltrimethylammonium bromide (CTAB) [4], sodium dodecyl sulfate (SDS) [5], urea [6], and guanidine hydrochloride [7], and optimized a cost- and time-efficient DNA extraction protocol for maize endosperm samples.
Materials and reagents
Biological materials
1. Maize kernels (seeds)
Reagents
1. GoTaq GreenMaster Mix 2× (Promega, catalog number: M7123)
2. Dimethyl sulfoxide (DMSO) (Promega, CAS 67-68-5)
3. Betaine (Promega, CAS 107-43-7)
Laboratory supplies
1. 2.2 mL 96-well sample collection plates (n+1 plates for n sets of 96 tissue samples; see step A1) (VWR, catalog number 43001-0020)
2. 1.2 mL sampling tubes (96 for one set of 96 tissue samples, if using SDS or filter plate DNA extraction) (VWR, catalog number 83009-678)
3. 2.0 mL safe-lock microcentrifuge tubes (96 for one set of 96 tissue samples, if using CTAB DNA extraction) (Eppendorf, catalog number 022363352)
4. Labeling tape
Equipment
1. Forceps (VWR, catalog number 82027-446)
2. Pet nail clipping pliers (Resco, catalog number: PF0728); see General Note 1
3. Mini ice cube tray, used for seed storage, 135 cubes per tray 0.5 inch size (e.g., Source 1 or Source 2).
Software
1. Tissue Sample Plate Mapper software; available as a Google sheet plug-in (Google Workspace
Marketplace, plug-in number: 115639436496)
2. Excel or other spreadsheets may be useful for additional notetaking
3. Primer3 software (https://bioinfo.ut.ee/primer3-0.4.0/)
4. MaizeGDB (http://www.maizegdb.org) for primer blast analysis
Procedure
A. Labeling seed trays and collection plates
1. During seed chipping, two 96-well sample collection plates are needed. One plate (the tube holder; Figure 1A) holds one sampling tube at a time to aid tissue collection during chipping.
2. The second 96-well sample collection plate (for tissue storage; Figure 1A) stores up to 96 accumulated sampling tubes containing chipped tissue. Optionally, prepare the tissue storage plate by printing a plate map using the tissue-sample plate mapper software, overlaying the map on a 96-well sample collection plate and securing it with clear tape (see General Note 4). Label this plate with a unique ID using tape and a marker.
3. One seed storage tray is used to hold chipped maize seeds until analysis and decision-making are completed. Lab tape may be used to block off superfluous sections so the tray holds 96 seeds in an 8 × 12 format (Figure 1A). Label this tray with the same unique ID to pair it with a tissue storage plate.
B. Maize seed chipping
See Video 1 for a demonstration of the complete chipping procedure.
1. On a large laboratory bench, place the tube holder plate, labeled tissue storage plate, and seed storage tray with the tube holder plate closest to you and the seed storage tray farthest from you (Figure 1A).
2. Place one sampling tube in one of the tube holder plate’s bottommost corner wells. To help contain debris produced in the chipping process, unused wells in the tube holder plate may be covered with masking tape (optional). The corner location for the sampling tube facilitates positioning the clipping pliers right above the sampling tube (Figure 2H, 2K).
3. Using the dominant hand, hold the clipping pliers' blade side up, with the thumb on the top, stationary handle, and the other fingers holding the lower handle with the spring attached that moves the blade.
4. With your other hand’s thumb and index finger, pick up and hold one maize seed by the tip cap with the embryo side facing toward the thumb (Figure 2H–2I).
5. While holding the chipper orifice directly and slightly above the sampling tube in the tube holder plate, hold the seed approximately perpendicular to the pliers, place its crown (Figure 2A) into the chipper hole, and gently chip the seed repeatedly to shave off fine tissue particles (Figures 2H–2J). For example, three to five passes of the chipping blade are usually required with B73 and Mo17 kernels.
Critical: Chip and shave the tissue finely (Figures 2B, 2D) because large chip particles (Figure 2F) grind poorly and barely yield extracted DNA (see General Note 2).
Critical: It is imperative to avoid damaging the embryo (Figure 2A) while chipping the endosperm. If necessary, secondary chips may be collected, e.g., from small to or rounded kernels like from the W22 inbred, by slightly tilting the seed at a 45° angle to the first plane of chipping. Then, chip and shave fine endosperm particles as described above for the crown, but instead chip from the side opposite the embryo (Figure 2K–2M). A combination of such primary and secondary chips also increases the proportion of endosperm tissue in the sample relative to maternal pericarp tissue.
Critical: Processing too much tissue may reduce DNA yield in the extraction step. Using the extraction methods tested, approximately 15–30 mg of tissue optimally yields approximately 5 μg of DNA at concentrations appropriate for use as PCR template.
6. If the SDS, filter plate, or urea method is used for DNA extraction (section C), then remove the sample tube with chipped tissue from the tube holder plate and place it into the tissue storage plate; then, place the chipped seed into the appropriate well in the seed tray. For tracking purposes, match the chipped seed’s position in the seed tray grid with the position of the sampling tube on the tissue storage plate grid. If CTAB is used for DNA extraction (section C), then transfer the chipped tissue into a labeled 2 mL microcentrifuge tube instead.
7. Clean the clipping pliers by wiping with a dry tissue and/or blowing off any chip residue. Application of compressed air while flexing the pliers’ handle may be used to ensure no chip dust carries over in or on the tool; more thorough cleaning of the chipping pliers may be performed periodically using 70% ethanol and compressed air.
8. Repeat steps B2–B7 with a new sampling tube and kernel until 96 or the desired number of samples have been collected.
9. When sampling is completed, seed storage trays may be stacked and placed in the sealed plastic tubs they are shipped in (see Equipment) and set aside for temporary storage. For a turnaround to sowing within six months, storage at room temperature and humidity had no apparent effects on germination frequency.
10. Proceed to DNA extraction.
Figure 1. Maize seed chipping setup and genotyping. A. Sampling includes clipping pliers, seeds, a 96-well plate acting as a tube holder plate, a 96-well tissue storage plate overlaid with a sampling map to accumulate chipped samples in tubes, and a seed storage tray. B. Once chipping is completed, chipped tissue samples are processed for DNA extraction, which is genotyped using PCR and an agarose gel and data analysis. C. With data tracking, seeds with the genotype of interest are selected for downstream applications.
Video 1. Chipping maize endosperm for DNA extraction
Figure 2. Maize kernels and the seed chipping process. A. Anatomy of a maize kernel in longitudinal section; peri: pericarp; endo: endosperm. B. An appropriate volume of seed chips in a collection tube. C. Intact, unchipped kernel. D, E. Properly chipped kernel shavings (D, green circle) removed from the crown of the kernel (E). F, G. Improperly chopped, large chunk(s) (F, red circle) removed from the crown (G). H–J. Seed chipping of the crown is done with the embryo facing you and the seed nearly perpendicular to the clipping pliers. K–M. Secondary chips, e.g., from round-shaped kernels, produced by holding the seed with the embryo facing upward and nearly parallel to the chipping pliers’ blade to shave chips from the back of the seed, i.e., at a 45° angle to the first chipping surface. Scale bars = 10 mm for panels H and K, scale bars = 2 mm for the remaining panels.
C. DNA extraction options
1. Guanidine-HCl filter plate extraction protocol (see Supplemental Protocol 1)
2. SDS-based DNA extraction protocol (see Supplemental Protocol 2)
3. Urea-based DNA extraction protocol (see Supplemental Protocol 3)
4. Modified CTAB DNA extraction protocol with steel-bead (https://bio-protocol.org/en/bpdetail?id=2906&type=0; [4])
D. PCR and genotyping
1. Design high-quality genotyping markers using the maize reference genome (http://www.maizegdb.org) and Primer3 (https://bioinfo.ut.ee/primer3-0.4.0/).
2. Set up a 13 μL PCR reaction composed of 2× GoTaq GreenMaster Mix (6 μL), DMSO (0.5 μL), forward and reverse primer mix (0.5 μL of 5 μM each), water (2.5 μL), and DNA (3 μL). For amplifying GC-rich regions, substitute the water in the PCR reaction with betaine (2.5 μL of 5 M).
3. Run standard PCR cycling parameters according to the primers used and product size. For instance, for Figure 4, the primers used were BZ316 (5'-ATCTGCATCCTGCGACGCAAC-3') and BZ317 (5'-GTCGGCGGTCTTTCTCGAGTC-3') and PCR conditions were 94 °C, 3 min (1 cycle); 94 °C, 30 s, 60 °C, 30 s, 72 °C, 1 min (34 cycles); 72 °C, 5 min (1 cycle); hold at 10 °C for forever. This reaction produced amplicon sizes of 555 bp (B73) and 416 bp (Mo17).
4. To genotype, run the PCR-amplified products with, e.g., 100 bp ladder marker on a 2% agarose gel in TAE buffer at 120 V for 20 min or longer depending on the product size.
5. Record the genotypes. If using the tissue-sample plate mapper software, then genotype calls may be entered in the ordered Sample List tab generated by the software.
6. After data analysis, use forceps to carefully pick chipped seeds with the genotype(s) of interest from the seed storage trays.
Data analysis
Seed chipping with nail clipping pliers provides safer, scalable, and better throughput than razor blade–based methods. Thirty to forty minutes are required to chip 96 maize seeds, depending on seed size and shape. Smaller or round seeds are challenging to hold and need more time to chip than larger ones. To examine the effects of seed chipping on germination, we planted chipped and unchipped seeds from the same two seed packets in three replications of 32 kernels in the greenhouse. One packet contained inbred B73 and the other contained inbred Mo17. Seeds were sown in prewet Metro-Mix 830 or similar in a standard maize greenhouse (16/8 h day/night photoperiod, ~27/18.5 °C, respectively) and covered with a 7-inch humidity dome to maintain moisture until emergence. Seedlings were watered sparingly, as needed. We scored germination per each kernel based on whether or not a coleoptile emerged above the soil line; we also measured vigor per each replicate as an aggregate (chipped or unchipped) based on any differences overall in growth rate or in appearance by a response to any unintended stresses that may have been present in their shared environmental conditions. We noticed that chipping did not impact the germination rate, but seedlings from chipped seeds initially grew more slowly (Figure 3; see General Note 3) while other observed responses occurred uniformly across both treatments, e.g., some yellowing occurred in all seedlings in the B73 experiment (Figure 3B).
Figure 3. Effect of seed chipping on germination and seedling growth across B73 and Mo17 maize inbreds. A. Germination frequency (y-axis, showing from 90% to 100%) of unchipped (blue) and chipped (red) seeds across two maize inbreds, Mo17 and B73 (x-axis). Each circle indicates the percentage germination from one replicate of 32 sown kernels. Each line indicates the mean for three replicates. B. Post-germination seedlings at 14 days to compare the growth of unchipped (left) and chipped seeds (right) for B73. C. Post-germination seedlings at 14 days to compare the growth of unchipped (left) and chipped seeds (right) for Mo17. Scale bars = 10 cm.
We compared the quality of DNA extracted from seeds to DNA extracted from leaves using several different DNA extraction methods. The results indicated that the quality of the DNA extracted from seeds and leaves was comparable using three of the four methods tested (Figure 4A). The exception was the urea-based method, where DNA extracted from seed chips performed poorly (Figure 4A). Regarding usability and high throughput, the extraction methods are user-friendly and high-throughput, except for CTAB. For instance, with CTAB extraction, we process a maximum of 48 samples at a time, whereas with other extraction methods, 384 samples can be processed at a time once chipping is complete. The guanidine-HCl with filter plate method produced the best-quality DNA but can include significant expenses for reagents and the filter plate. Thus, we conclude that for a seed chip sample type, the SDS extraction method optimizes the combination of usability, high throughput, and cost-effectiveness (e.g., figure 4B).
In maize seed-based genotyping, maternal pericarp tissue has led to incorrect genotype results. Gao et al. [2] reported that among the homozygotes within an F2 population, false heterozygous genotyping errors occurred at a rate of 3.8% due to maternal pericarp DNA when chipping was performed after soaking maize kernels in water. While pericarp “contamination” could be reduced or eliminated by carefully peeling pericarps away prior to chipping endosperm, that approach is labor intensive. To minimize pericarp contribution, we included secondary chips that enriched chip samples for endosperm, diluting the proportion of pericarp sampled. To quantify the significance of pericarp “contamination,” we compared genotype results in an F2 population from B73 × Mo17 by first seed-chipping 96 F2 kernels and then germinating and leaf-sampling their corresponding 96 F2 seedlings. DNA was prepared using the SDS method for both seed chip and leaf tissue and PCR was performed as above (Section D). For 88 of the 96 individuals, sufficient PCR product was detected to call genotypes for both the chip- and leaf-based methods. Among the 88 individuals, we focused on the homozygotes because in F2 seed-chip DNA, either homozygous genotype could theoretically be miscalled as heterozygous due to PCR amplification of DNA extracted from the heterozygous, maternal pericarp tissue. 47 individuals were homozygous, as called by the leaf-based method. Among their corresponding chip-DNA genotypes, 43 had the homozygous call, 4 could not be called due to no PCR product, and none were miscalled. Thus, we observed zero disparities between the seed and leaf DNA groups, and therefore found no evidence supporting pericarp contamination among the 43 homozygotes (e.g., see Figure 4B–4C).
Figure 4. PCR-based genotyping using DNA extracted from chipped seed endosperm and comparison with DNA from leaves. A. Comparison of seed-based and leaf-based DNA across four DNA extraction methods. B–C. Testing for pericarp-based genotyping miscalls by comparing corresponding chipped (B) seed- and (C) leaf-based DNA from the same 96 individuals; representative data from 48 individuals (1–48, yellow numbers) are shown. Homozygotes show upper band alone (e.g., individual 7) or lower band alone (e.g., individual 29); heterozygotes show both bands (e.g., individual 6). M: marker DNA; New England Biolabs 100 bp DNA ladder; intense bands are 500 bp and 1,000 bp.
Validation of protocol
This protocol was used in the authors’ research over the last year to chip and genotype over 1,300 kernels and then selectively germinate the desired class(es). We used SDS-based DNA extraction and genotyped at six different loci for which we had previously developed PCR assays using DNA extracted from leaves. All six assays worked well with DNA extracted from chipped endosperm and the results depicted in Figure 4B are highly representative; genotype calls were returned for upward of 90% of the chipped kernels. Chipped kernels generally germinated at the same frequency as unchipped ones.
General notes and troubleshooting
General notes
1. Plastic and metal nail clipping pliers were tested (Figure 5); we recommend the metal pliers because they are durable and the blade can be removed for sharpening or replacement.
Figure 5. Nail clipping pliers tested. Clippers for seed chipping were either metal with an exchangeable blade (A, preferred) or plastic with a fixed blade (B).
2. Sufficient chipped seed tissue (15–30 mg) should be collected for adequate DNA for one to several genotyping reactions. Within that constraint, keep the chipped tissue pieces as small as possible because larger particles are more challenging to grind in the DNA extraction step. During the chipping process, chip more endosperm to dilute and minimize the amount of pericarp sampled.
3. Seedlings from chipped seeds grew slower, perhaps due to reduced mobilization of nutrients from the reduced volume of aleurone and endosperm. Thus, we also recommend not chipping more endosperm than required for adequate DNA yield (see note 2). Moreover, chipped seeds are susceptible to mold infection, especially if germinated in paper towels. Thus, treating or disinfecting the seed before planting may be warranted.
4. Additional details on installing and using the plate mapping software as a Google sheet plug-in are available through the Vollbrecht lab website (https://faculty.sites.iastate.edu/vollbrec/tissue-sample-plate-mapper). When printing plate maps using the software, navigate to the Plate labels for printing tab and execute the print command with these settings: Print current sheet; Paper size, letter; Landscape orientation; Scale, normal (100%); Margins, normal; Formatting, show gridlines and show notes; Page order, over then down; Alignment horizontal, center; Alignment vertical, center; Headers and Footers selections according to your preferences.
5. Trays with chipped seeds may be stored at room temperature for up to six months. Cool (to prevent damage from pests such as mice or moths) and low-humidity conditions (e.g., 8 °C, ~25% relative humidity) would be optimal for maize, especially for long-term storage [8]. A refrigerator or cold room that is not humidity-controlled would result in high humidity levels (~ 100% relative humidity) and would be detrimental to viability over time; it is less desirable than storing at room-temperature conditions. We have not investigated how chipping affects overall seed storage longevity.
Supplementary information
The following supporting information can be downloaded here
Supplemental Protocol 1
Supplemental Protocol 2
Supplemental Protocol 3
Acknowledgments
The tissue-sample plate mapper software was developed by Takao Shibamoto and Kokulapalan Wimalanathan with support from the National Science Foundation NSF-IOS grant 1238202 to E. Vollbrecht. Some figures were generated with BioRender.com. This research was also supported by funding from the Iowa State University Crop Bioengineering Center.
Competing interests
The authors declare no competing interests.
References
Xu, Y. and Crouch, J. H. (2008). Marker‐Assisted Selection in Plant Breeding: From Publications to Practice. Crop Sci. 48(2): 391–407. https://doi.org/10.2135/cropsci2007.04.0191
Gao, S., Martinez, C., Skinner, D. J., Krivanek, A. F., Crouch, J. H. and Xu, Y. (2008). Development of a seed DNA-based genotyping system for marker-assisted selection in maize. Mol Breed. 22(3): 477–494. https://doi.org/10.1007/s11032-008-9192-4
Mills, A., Allsman, L., Leon, S. and Rasmussen, C. (2020). Using Seed Chipping to Genotype Maize Kernels. Bio Protoc. 10(6): e3553. https://doi.org/10.21769/bioprotoc.3553
Yi, S., Jin, W., Yuan, Y. and Fang, Y. (2018). An Optimized CTAB Method for Genomic DNA Extraction from Freshly-picked Pinnae of Fern, Adiantum capillus-veneris L. Bio Protoc. 8(13): e2906. https://doi.org/10.21769/bioprotoc.2906
Edwards, K., Johnstone, C. and Thompson, C. (1991). A simple and rapid method for the preparation of plant genomic DNA for PCR analysis. Nucleic Acids Res. 19(6): 1349–1349. https://doi.org/10.1093/nar/19.6.1349
Leach, K. A., McSteen, P. C. and Braun, D. M. (2016). Genomic DNA Isolation from Maize (Zea mays) Leaves Using a Simple, High‐Throughput Protocol. Curr Protoc plant Biol. 1(1): 15–27. https://doi.org/10.1002/cppb.20000
Gao, H., Smith, J., Yang, M., Jones, S., Djukanovic, V., Nicholson, M. G., West, A., Bidney, D., Falco, S. C., Jantz, D., et al. (2010). Heritable targeted mutagenesis in maize using a designed endonuclease. Plant J. 61(1): 176–187. https://doi.org/10.1111/j.1365-313x.2009.04041.x
Guzzon, F., Gianella, M., Velazquez Juarez, J. A., Sanchez Cano, C. and Costich, D. E. (2021). Seed longevity of maize conserved under germplasm bank conditions for up to 60 years. Ann Bot. 127(6): 775–785. https://doi.org/10.1093/aob/mcab009
Article Information
Publication history
Received: Apr 4, 2024
Accepted: Dec 5, 2024
Available online: Jan 9, 2025
Published: Feb 5, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Plant Science > Plant molecular biology > Genetic analysis
Molecular Biology > DNA > DNA extraction
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5,201 | https://bio-protocol.org/en/bpdetail?id=5201&type=0 | # Bio-Protocol Content
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This is an In Press version of the protocol that has not yet been assigned to an issue.
Peer-reviewed
Development and Application of MLB Human Astrovirus Reverse Genetics Clones and Replicons
HA Hashim Ali
DN David Noyvert
Valeria Lulla
In Press, Available online: Jan 19, 2025
DOI: 10.21769/BioProtoc.5201 Views: 64
Reviewed by: Jibin SadasivanVamseedhar Rayaprolu Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in PLOS Biology Jul 2023
Abstract
Human astroviruses pose a significant public health threat, especially to children, the elderly, and immunocompromised individuals. Nevertheless, these viruses remain largely understudied, with no approved antivirals or vaccines. This protocol focuses on leveraging reverse genetics (RG) and replicon systems to unravel the biology of MLB genotypes, a key group of neurotropic astroviruses. Using reverse genetics and replicon systems, we identified critical genetic deletions linked to viral attenuation and neurotropism, pushing forward vaccine development. We also uncovered novel replication mechanisms involving ER membrane interactions, opening doors to new antiviral targets. Reverse genetics and replicon systems are essential for advancing our understanding of astrovirus biology, identifying virulence factors, and developing effective treatments and vaccines to combat their growing public health impact.
Key features
• Provides a basic understanding of the molecular biology of MLB astroviruses, aiding in addressing open questions related to virus evolution, replication, and pathogenesis.
• Facilitates the development of novel therapeutics and vaccines.
• Enables rapid testing of antiviral drugs against MLB astroviruses.
Keywords: Human astrovirus Reverse genetics Replicon Virus rescue Virus passaging Replication Replicon assay and immunodetection
Graphical overview
Background
Human astroviruses (HAstVs) are small, positive-sense RNA viruses primarily associated with gastroenteritis, causing symptoms such as diarrhea, vomiting, and abdominal cramps. These viruses significantly impact public health, particularly affecting young children, the elderly, and immunocompromised individuals, accounting for up to 9% of pediatric gastroenteritis cases globally. There are three major groups of human astroviruses: classical HAstVs, newly emerging HAstV-MLB (Melbourne), and HAstV-VA/HMO (Virginia/human-mink-ovine-like) astroviruses. These viruses are globally prevalent, infect a wide range of species, and demonstrate the capacity for rapid evolution and adaptation to different hosts. Despite their medical importance, many human astroviruses remain understudied due to the lack of essential molecular tools, such as reverse genetics (RG) infectious clones and replicon systems. In addition to causing gastrointestinal disease, certain astroviruses, particularly the MLB genogroup, have been implicated in severe neurological complications, such as encephalitis and meningitis. The MLB genogroup includes distinct viral strains: MLB1, MLB2, and MLB3. Among these, MLB1 is the most prevalent genotype, followed by MLB2 and MLB3. All MLB stains share similarities, including genomic organization and neurotropic potential. Recent studies [1] have developed critical molecular biology tools, such as reverse genetics systems, replicon systems, and immunodetection assays, to investigate the key stages of MLB astrovirus infection. Additionally, novel replication mechanisms were recently uncovered using the MLB RG and replicon systems [2]. These findings revealed interactions between endoplasmic reticulum (ER) membranes and replication factories located in close proximity to the nuclear periphery in a nsP1-dependent manner, providing new insights and potential targets for antiviral therapies.
Materials and reagents
Biological materials
1. Huh7.5.1 cells (obtained from Apath, Brooklyn, New York, United States of America)
2. MLB1 RG infectious virus stocks (from this study, store at -70 °C)
3. MLB2 RG infectious virus stocks (from this study, store at -70 °C)
4. MLB1 replicon based on the reverse genetics of MLB1 (from this study, -20 °C)
5. MLB2 replicon based on the reverse genetics of MLB2 (from this study, -20 °C)
Reagents
1. Dulbecco’s modified Eagle’s medium (DMEM) (PAN Biotech, catalog number: P04-03600)
2. Fetal bovine serum (FBS) (PAN Biotech, catalog number: P40-37500)
3. Penicillin-streptomycin (10,000 U/mL) (Life Technologies, catalog number: 15140-122)
4. 200 mM L-Glutamine (Gibco, catalog number: 25030081)
5. Phosphate buffered saline (PBS) (Life Technologies, catalog number: 14190-144)
6. 1 M HEPES buffer, pH range 7.2–7.5 (Life Technologies, catalog number: 15630-080)
7. 0.25% Trypsin-EDTA (Gibco, catalog number: 25200056)
8. Bovine serum albumin (BSA) (PAN Biotech, catalog number: P06-1395500)
9. Direct-zolTM RNA MiniPrep Plus (Zymo Research, catalog number: R2072)
10. PhusionTM Plus DNA polymerase (Thermo Fisher Scientific, catalog number: F630S)
11. XhoI (Thermo Fisher Scientific, catalog number: FD0695)
12. OptiMEM I (Gibco, catalog number: 31985070)
13. RNA Clean-Up and Concentrator kit (Zymo Research, catalog number: R1016)
14. RNaseOUT (Invitrogen, catalog number: 10777019)
15. Paraformaldehyde solution (PFA), 4% in PBS (Thermo Fisher Scientific, catalog number: 30525-89-4)
16. Βeta-mercaptoethanol (Sigma, catalog number: M3148)
17. T7 mMESSAGE mMACHINE Transcription kit (Thermo Fisher Scientific, catalog number: AM1344)
18. T7 Firefly luciferase plasmid (Promega, catalog number: L482A)
19. QIAquick Gel Extraction kit (Qiagen, catalog number: 28704)
20. QIAquick PCR Purification Kit (Qiagen, catalog number: 28104)
21. Dual-Luciferase Reporter Assay System (Promega, catalog number: E1980)
22. Anti-CP antibody (custom-made rabbit polyclonal antibody)
23. Anti-tubulin antibody (Abcam, mouse monoclonal antibody, catalog number: ab6160)
24. IRDye 800 CW goat anti-rabbit secondary antibody (LI-COR Biotechnology, catalog number: 926-32211)
25. IRDye 680 RD goat anti-mouse IgG secondary antibody (LI-COR Biotechnology, catalog number: 926-68070)
26. Anti-dsRNA antibody (anti-dsRNA IgG2a) (Scicons J2, catalog number: 10010500)
27. Goat anti-rabbit IgG (H+L) secondary antibody Alexa 488-conjugated (Thermo Fisher Scientific, catalog number: A-11008)
28. Goat anti-mouse IgG (H+L) secondary antibody 594-conjugated (Thermo Fisher Scientific, catalog number: A-11005)
29. Nitrocellulose membrane, 0.2 μm (Thermo Fisher Scientific, catalog number: 15209874)
30. 8% Sodium azide solution (Thistle Scientific, catalog number: SBL-40-2008-01)
31. MEM non-essential amino acid solution (Sigma, catalog number: M7145)
32. GibcoTM VP-SFM media (Fisher Scientific, catalog number: 10593273)
33. Normal goat serum (Abcam, catalog number: ab7481)
34. DNase I (Zymo Research, catalog number: E1011)
35. Lipofectamine 2000 (Thermo Fisher Scientific, catalog number: 11668019)
36. QIAGEN Plasmid Plus Midi Sample kit (Qiagen, catalog number: 12941)
37. Agarose (Sigma-Aldrich, catalog number: A9539)
38. Tween-20 (Sigma-Aldrich, catalog number: P1379)
39. PIPES (Sigma-Aldrich, catalog number: P6757)
40. EGTA (Sigma-Aldrich, catalog number: 03777)
41. MgCl2 (Sigma-Aldrich, catalog number: 31413)
42. NaOH (Sigma-Aldrich, catalog number: 06203)
43. Glycerol (Sigma-Aldrich, catalog number: G5516)
44. Triton X-100 (Thermo Fisher Scientific, catalog number: HFH10)
45. Tris HCl (Duchefa Biochemie, catalog number: T1513)
46. SDS (Sigma-Aldrich, catalog number: 75746)
47. pAVIC plasmid [3]
48. Bromophenol blue (Sigma-Aldrich, catalog number: B0126)
49. Dried skimmed milk powder (Marvel, multiple suppliers, grocery stores)
50. Formaldehyde, methanol free (Polysciences, catalog number: 04018-1)
51. Hoechst stain (Thermo Fisher Scientific, catalog number: 62249)
52. Wheat germ agglutinin (WGA) (Thermo Fisher Scientific, catalog number: W11261)
53. Perasafe (Medisave, catalog number: UN068)
54. Chemgene (Thermo Fisher Scientific, catalog number: SKU075A)
55. Qiagen AVL lysis buffer (Qiagen, catalog number: 19073)
56. Zymo-Spin IC columns (Zymo Research, catalog number: C1004-50)
57. Invitrogen SuperScript III Reverse Transcriptase (Thermo Fisher Scientific, catalog number: 18080044)
58. RNase-free water (Qiagen, catalog number: 129112)
59. 10% Chloros (sodium hypochlorite) (Thermo Fisher Scientific, catalog number: 219255000)
Solutions
1. Complete media (DMEM-GHAA-10% FBS) (see Recipes)
2. DMEM-GHAA-5% FBS media (see Recipes)
3. Infection media (DMEM-GHAA+0.2% BSA) (see Recipes)
4. DMEM-GH-2% FBS media (see Recipes)
5. DMEM-GH media (see Recipes)
6. Transfection media (15 mL)
7. Serum-free media (DMEM-GHAA) (see Recipes)
8. VPSF-GHAA media (see Recipes)
9. Antibody solution (see Recipes)
10. 2× PHEM buffer (see Recipes)
11. 2× Loading buffer + beta mercaptoethanol (2× LB-βME) (see Recipes)
12. 0.08% Trypsin-EDTA solution (see Recipes)
13. 1% Perasafe solution (see Recipes)
14. 5% Chemgene solution (see Recipes)
15. 0.1% PBST solution (see Recipes)
Recipes
1. Complete media (DMEM-GHAA-10% FBS) (500 mL)
Reagent Final concentration Quantity or Volume
DMEM n/a 429 mL
100% FBS 10% 50 mL
1 M HEPES 20 mM 10 mL
Penicillin and streptomycin 20 Units/mL 1 mL
100× MEM non-essential amino acids n/a 5 mL
200 mM L-glutamine 2 mM 5 mL
Total (optional) n/a 500 mL
2. DMEM-GHAA-5% FBS media (500 mL)
Reagent Final concentration Quantity or Volume
DMEM n/a 454 mL
100% FBS 5% 25 mL
200 mM L-glutamine 2 mM 5 mL
1 M HEPES 20 mM 10 mL
Penicillin and streptomycin 20 Units/mL 1 mL
100× MEM non-essential amino acid n/a 5 mL
Total (optional) 500 mL
3. Infection media (DMEM-GHAA+0.2% BSA) (50 mL)
Reagent Final concentration Quantity or Volume
DMEM n/a 46.9 mL
200 mM L-glutamine 2 mM 0.5 mL
1 M HEPES 20 mM 1 mL
Penicillin and streptomycin 20 Units/mL 0.1 mL
100× MEM non-essential amino acid n/a 0.5 mL
10% BSA 0.2% 1.0 mL
Total (optional) n/a 50 mL
4. DMEM-GH-2% FBS media (500 mL)
Reagent Final concentration Quantity or Volume
DMEM n/a 475 mL
100% FBS 2% 10 mL
1 M HEPES 20 mM 10 mL
200 mM L-glutamine 2 mM 5 mL
Total (optional) n/a 500 mL
5. DMEM-GH media (500 mL)
Reagent Final concentration Quantity or Volume
DMEM n/a 485 mL
200 mM L-glutamine 2 mM 5 mL
1 M HEPES 20 mM 10 mL
Total (optional) n/a 500 mL
6. Transfection media (15 mL)
Reagent Final concentration Quantity or Volume
OptiMEM n/a 15 mL
RNaseOUT 40 Units/mL 15 μL
Total (optional) n/a 15 mL
7. Serum-free media (DMEM-GHAA) (500 mL)
Reagent Final concentration Quantity or Volume
DMEM n/a 479 mL
1 M HEPES 20 mM 10 mL
Penicillin and streptomycin 20 Units/mL 1 mL
100× MEM non-essential amino acids n/a 5 mL
200 mM L-glutamine 2 mM 5 mL
Total (optional) n/a 500 mL
8. VPSF-GHAA media (50 mL)
Reagent Final concentration Quantity or Volume
VP-SFM media n/a 47.9 mL
1 M HEPES 20 mM 1 mL
Penicillin and streptomycin 20 Units/mL 0.1 mL
100× MEM non-essential amino acids n/a 0.5 mL
200 mM L-glutamine 2 mM 0.5 mL
Total (optional) n/a 50 mL
9. Antibody solution (100 mL)
Reagent Final concentration Quantity or Volume
PBS 1× 100 mL
100% goat serum 0.5% 0.5 mL
8% sodium azide 0.02% 0.25 mL
Total (optional) n/a 100 mL
10. 2× PHEM buffer (500 mL)
Reagent Final concentration Quantity or Volume
PIPES 120 mM 72.56 g
HEPES 50 mM 23.84 g
EGTA 20 mM 15.2 g
2 M MgCl2 4 mM 1 mL
H2O n/a see note*
Total (optional) n/a 500 mL
*First dissolve PIPES in 400 mL of water and then add the other reagents. Adjust pH to 7.2 with 5 M NaOH or KOH.
11. 2× loading buffer + beta-mercaptoethanol (2× LB-βME) (10 mL)
Reagent Final concentration Quantity or Volume
1 M Tris HCl pH 6.8 125 mM 1.25 mL
10% SDS 4% 4 mL
1% Bromophenol blue 0.04% 0.4 mL
100% glycerol 20% 2 mL
Beta-mercaptoethanol 10% 1 mL
Then, add water to make a final volume of 10 mL.
12. 0.08% Trypsin-EDTA solution
Add 2.5 mL of 0.25% trypsin-EDTA in 7.5 mL of PBS.
13. 1% Perasafe solution
1 g of Perasafe powder in 100 mL of water.
14. 5% Chemgene solution
Add 50 mL of Chemgene in 1,000 mL of water.
15. 0.1% PBST solution
Add 1 mL of Tween-20 in 1,000 mL of PBS.
Laboratory supplies
1. Pipette tips (Thermo Fisher Scientific, catalog numbers: 02-707-426, 02-707-403, 02-707-438)
2. 1.5 mL microfuge tubes (STARLAB, catalog number: 51615-5500)
3. 15 mL Falcon tubes (Corning, catalog number: 430052)
4. 50 mL Falcon tubes (Corning, catalog number: 430829)
5. T175 tissue culture flasks (TPP® tissue culture flasks) (Sigma, catalog number: Z707562)
6. 6-multiwell polystyrene culture plates (TPP® tissue culture plates) (Sigma, catalog number: Z707767)
7. 12-multiwell polystyrene culture plates (TPP® tissue culture plates) (Sigma, catalog number: Z707775)
8. IBIDI 8-well chambered slides (Ibidi GmbH, catalog number: 80807-90)
9. 96-well flat-bottom plate (TPP, catalog number: Z707902)
10. U-bottom 96-well plate (TPP, catalog number: Z707899)
11. 0.2 μM filters (Appleton, catalog number: ACF141)
12. Screw-cap tubes (Axigen, catalog number: SCT-150-A-S)
Equipment
1. Biosafety level 2 culture cabinet (Wolflabs, model: BioMAT 2 BM21800R)
2. -70 °C freezer (Eppendorf, catalog number: F660320001)
3. Vortex (Heidolph, model: Reax Top)
4. Spray bottle with 70% ethanol (Sigma-Aldrich, catalog number: 32221)
5. Pipettes: 1,000 μL (StarLab, catalog number: S1111-6001)
6. Pipettes: 200 μL (StarLab, catalog number: S1111-0000)
7. Pipettes: 10 μL (StarLab, catalog number: S1111-3000)
8. Pipettes: 2 μL (StarLab, catalog number: S1111-3000)
9. 25 mL pipettes (Corning, catalog number: 760 180)
10. 10 mL pipettes (Corning, catalog number: 4488)
11. CO2 incubator (PHCBI, catalog number: MCO-230AIC)
12. Hemocytometer (Logos BioSystems, catalog number: L40002)
13. Cell counting slides (Logos BioSystems, catalog number: L12002)
14. Rocking platform shaker (Cole-ParmerTM, catalog number: WZ-51900-30)
15. Phase-contrast inverted microscope (Nikon TMS Inverted Microscope)
16. 4 °C refrigerator (LabCold, catalog number: RLPR0517)
17. Autoclave (BMM Weston, custom-made)
18. Bio-Rad Gene Pulser Xcell system (Bio-Rad, catalog number: 1652661)
19. Electroporation cuvettes (Bio-Rad, catalog number: 1652091)
20. Water bath (Clifton, catalog number: ZT1250337S)
21. LI-COR ODYSSEY CLx imager (LicorBio, catalog number: 9140-09)
22. Leica SP5 confocal microscope (Leica Microsystems, catalog number: Leica DMI6000 CS)
23. Trans-Blot Turbo transfer system (Bio-Rad, catalog number: 1704150EDU)
24. SDS-PAGE running assembly (Bio-Rad, catalog number: 1658001FC)
25. GloMax® Navigator microplate luminometer (Promega, catalog number: GM2010)
26. NanoDropTM 2000 spectrophotometer (Thermo Fisher Scientific, catalog number: ND2000)
Software and datasets
1. ImageJ 1.47v (10.2)
2. Prism v8.2.2 (GraphPad, July 2019)
Procedure
CRITICAL: All experiments with MLB RG viruses (i.e., all pre-fixation and pre-lysis steps) should be performed inside a biosafety level 2 (BSL2/CL2) tissue culture laboratory according to the country and institution regulations and required permits regarding handling and storage of human astroviruses.
Part I: Production of infectious recombinant MLB1 and MLB2 astrovirus stocks
A. Engineering MLB reverse genetics
1. To construct RG clones for MLB1 and MLB2, specific primers are designed using the 5' and 3'-terminal consensus sequences (Table 1).
Table 1. List of primers
Primers Sequences (5' to 3')
1 MLB1 forward GAGTAATACGACTCACTATAGCCAAGAGTGGTGGTATGGCTG
2 MLB1 reverse CCATACATTTATGCTGGAAGAAAAAAAGC
3 MLB2 forward GAGTAATACGACTCACTATAGCCAAGAGTGGTAGGATGGCTGTG
4 MLB2 reverse CCTCTAAATCTACCTGATTAGAAAAAAAAAGATAAAATTTTATTTGTC
2. The viral genomic RNAs are extracted from clinical isolates (GenBank accession number MLB1-MK089434 and MLB2-MK089435) using Qiagen AVL lysis buffer and Zymo spin RNA purification columns.
3. Elute viral genomic RNAs in 30 μL of RNase-free water and quantify using a Nanodrop.
4. For cDNA synthesis, use 11 μL of viral RNA (25–100 ng) using Superscript III reverse transcriptase (Table 2).
Table 2. cDNA synthesis reaction mixture 1
Component Volume (μL)
1 viral RNA 11 μL (25 ng to 100 ng)
2 dNTPs 1.0 μL (10 mM)
3 Virus-specific reverse primer 1.0 μL (100 μM)
Total volume 13 μL
5. Incubate room temperature (RT) reaction at 65 °C for 5 min and cool down to 4 °C.
6. Add 7 μL of reverse transcriptase mix to each PCR reaction (4 μL of 5× first strand buffer + 1 μL of 100 mM DTT (Dithiothreitol) + 1 μL of RNaseOUT +1 μL of SuperScript III Reverse Transcriptase enzyme).
7. Incubate at 55 °C for 60 min, followed by incubation at 70 °C for 15 min and at 10 °C on hold.
8. Full-length genomes of MLB1 and MLB2 are amplified by PCR using Phusion High-Fidelity DNA polymerase and virus-specific primer sets (Table 3).
Table 3. PCR amplification reaction
Component Volume (μL)
1 cDNA (template), 25 ng to 100 ng 1.0 μL
2 5× HF buffer 10.0 μL
3 10 mM dNTPs 1.0 μL
4 Virus-specific forward primer, 100 μM 0.3 μL
5 Virus-specific reverse primer, 100 μM 0.3 μL
6 Phusion DNA polymerase 0.5 μL
7 DMSO 1.0 μL
8 Water 36.0 μL
Total volume 50 μL
9. Perform PCR using the following steps: (step 1) 98 °C for 1 min; (step 2, 35 cycles): 98 °C for 10 s, 57 °C for 10 s, 72 °C for 3 min; (step 3): 72 °C for 5 min; and hold at 10 °C.
10. Separate amplified viral genomes on 0.7% agarose gel, purify using Zymo column, and sequence with specific primers.
11. The amplified viral genomes of MLB1 and MLB2 are cloned into the plasmid pAVIC [3] by replacing HAstV1 genome with MLB1 or MLB2 under T7 promoter using a single-step ligation-independent cloning method.
12. See plasmid maps of reverse genetics clones of MLB1 and MLB2 in Figure 1.
Figure 1. Plasmid maps of reverse genetics (RG) clones of MLB1 and MLB2, generated using SnapGene. The map on the left represents MLB1, while the map on the right represents MLB2.
13. RG plasmids were transformed into XL1Blue competent cells using a standard protocol. Single positive colonies were selected to further amplify bacterial cultures, followed by plasmid purification using QIAGEN Plasmid Plus Midi kit.
B. Linearization and in vitro synthesis of T7 RNA transcripts
1. To linearize, digest the MLB RG clones with the XhoI restriction enzyme (as shown in Table 4) and their corresponding GNN mutants (RdRp knockout as negative control).
Table 4. Restriction digestion reaction
Component Volume (μL)
1 Plasmids: MLB1/MLB2/GNN (1 μg/μL) 5.0 μL
2 XhoI (10 U/μL) 1.0 μL
3 10× FD buffer 5.0 μL
4 MilliQ water 39.0 μL
Total volume 50 μL
2. Incubate digestion reactions at 37 °C either in a water bath or incubator for 2 h.
3. Purify the linearized plasmids using a PCR Clean-Up kit using standard manufacturer’s instructions and elute in 20 μL of nuclease-free water or elution buffer. Quantify the concentrations of the linearized plasmids using a Nanodrop.
4. Synthesize T7 RNA transcripts using the T7 mMESSAGE mMACHINE Transcription kit (as shown in Table 5).
Table 5. In vitro synthesis of MLB RG T7 RNA transcripts
Component Volumes
1 Linear plasmids (MLB1/MLB2/GNN) (150–400 ng) 3.0 μL
2 2× NTP/CAP 5.0 μL
3 10× transcription buffer 1.0 μL
4 T7 Enzyme mixture 1.0 μL
Total reaction volume 10 μL
5. Incubate the T7 RNA reactions at 37 °C for 1 h (Note: No DNaseI treatment). To evaluate the quality of the T7 RNA samples, check them by loading 0.5 μL of RNA on a 1% agarose gel before electroporation into Huh7.5.1 cells. At this point, the T7 RNA samples can be directly used for electroporation or stored at -70 °C.
C. Electroporation of T7 RNA transcripts
1. First, check the Huh7.5.1 cells’ quality and confluency; they should be 80%–90% confluent (T175 flask is enough for 3–4 electroporations in the 6-multiwell plate format) before plating. Using cells from overconfluent flasks would impact the electroporation efficiency of T7-transcribed RNA(s). Prechill the required number of electroporation cuvettes in closed plastic bags (number of samples + mock) at 4 °C to avoid moisture condensation.
2. Prewarm complete media and 0.08% trypsin-EDTA solution at 37 °C.
3. Discard the culture media from the flask and rinse Huh7.5.1 cells with 10 mL of 1× PBS.
4. Add 15 mL of 1× PBS and incubate for at least 3–5 min (to completely remove the FBS-containing media).
5. Add 5 mL of prewarmed 0.08% trypsin-EDTA, seal the flask completely, and then gently rotate the flask to ensure that trypsin is equally distributed to the monolayer of Huh7.5.1 cells. Incubate at room temperature for 2–3 min to trypsinize the cells.
6. Add 10 mL of complete media to neutralize trypsin, resuspend cells properly using a pipette, and collect cells into a 50 mL Falcon tube.
Note: Cells should not be over-trypsinized as it can reduce electroporation efficiency.
7. Centrifuge cells at 400× g for 5 min with no brake (it will take 10–15 min to stop). Discard the supernatant and carefully remove any leftover media using a 1 mL pipette.
8. Add 45 mL of 1× PBS to the cell pellets. If the cells are in more than one Falcon tube, combine them into one Falcon tube for washing. At this point, count the cells using a hemocytometer; ~5 × 105 cells are enough for one electroporation. Take the required number of cells for electroporation and centrifuge the cells at 400× g for 5 min with no brake function.
9. Discard PBS and resuspend again cells in PBS (800 μL of PBS per electroporation).
10. Proceed immediately to electroporation using Bio-Rad Gene Pulser Xcell system. Start with mock cell electroporation (use the same volume of 1× transcription buffer), then proceed with viral T7-transcribed RNA(s) from Table 5.
11. Add 750 μL of cells and 10 μL (~20 μg) of T7-transcribed RNA(s) into the cuvette and mix well.
12. Set the electroporator to the exponential protocol, 800 V, 25 μF, 4 mm cuvette. Pulse once; the expected time constant should be 0.3–0.5 ms. Mix the cells in the closed cuvette by tapping, then repeat the pulse with the same expected time constant.
13. Quickly add 500 μL of DMEM-GHAA-5% FBS media to the cuvette, mix, and transfer the cells to 1.5 mL microcentrifuge tubes. The same procedure is to be followed with other samples.
14. Centrifuge cells at 400× g for 5 min, discard the media, and add 1 mL of fresh DMEM-GHAA-5% FBS media. Seed the cells into 6-multiwell plates in 1 mL of total volume.
15. Three to five hours post seeding, change the media to 1.2 mL of DMEM-GHAA-5% FBS to remove dead cells and non-electroporated RNA transcripts.
16. Carefully wash cells after 24 h of post-electroporation with 1 mL of PBS.
17. Add 1.2 mL of VPSF-GHAA media and incubate electroporated cells for 24 h (MLB2) or 48–72 h (MLB1). Virus replication induces a cytopathic effect (CPE), which can be visualized under the microscope or quantified after fixing by staining nuclei and cells with Hoechst and WGA-FITC (Figure 2).
Figure 2. Representative images showing the MLB2 and MLB1 virus-induced CPE in Huh7.5.1 cells
18. To ensure proper cell lysis, freeze the plate at -70 °C for at least 1 h, then thaw at room temperature. Collect both cells and the supernatant in the 1.5 mL microcentrifuge tubes.
19. Clarify viral supernatants by centrifugation at 7,000× g for 5 min, and then filter supernatants through a 0.2 μm filter.
20. Carefully transfer viral supernatants to a screw-cap tube and supplement with 5% glycerol and 0.2% BSA (final concentrations). These virus stocks are labeled as passage 0 (P0).
21. Generate P1, P2, etc., virus stocks by infecting cells with P0 virus using MOI 0.1 (Figure 3).
Figure 3. Method for a plasmid-derived reverse genetics (RG) system for MLB1 and MLB2. MLB cDNAs contain the entire genome flanked by the T7 promoter and XhoI linearization site. Huh7.5.1 cells were electroporated with full-genome T7 transcripts; the collected virus was used for serial passages (P1 to P10) in the same cell line (adapted from Ali et al. [1]).
D. Titration of virus stocks using immunofluorescence-based detection
1. Seed Huh7.5.1 cells (2.5–3 × 104 cells per well) on 96-well flat-bottom plates 24 h before infection.
Note: The cells should be 80%–90% confluent at the time of infection.
2. Prepare 10-fold serial dilutions of virus stocks in the U-bottom 96-well plate. For 10-fold dilutions, use 135 μL of infection media and 15 μL of virus stock.
3. An example of a 96-well plate scheme used for titration of MLB1 virus stocks is given in Table 6.
Table 6. Preparation of virus dilutions
Dilution 1 2 3 4 5 6 7 8 9 10 11 12
10-1 MLB1 P1 MLB1 P2 MLB1 P3 MLB1 P4 MLB1 P5 MLB1 P6 MLB1 P7 MLB1 P8 MLB1 P9 MLB1 P10 Mock MLB1
GNN
10-2 ↓ ↓ ↓ ↓ ↓ ↓ ↓ ↓ ↓ ↓ ↓ ↓
10-3 ↓ ↓ ↓ ↓ ↓ ↓ ↓ ↓ ↓ ↓ ↓ ↓
10-4 ↓ ↓ ↓ ↓ ↓ ↓ ↓ ↓ ↓ ↓ ↓ ↓
4. Discard the culture media from overnight cultured cells using a multichannel pipette or by inverting the plate against 5–10 layers of paper towels.
5. Infect monolayers of cells with 100 μL of diluted viruses.
Note: Start from the lowest dilutions first, using the same multichannel pipette for the entire plate.
6. Incubate the infected cells in the incubator (5% CO2, 37 °C) for 20–28 h (MLB2) and 40–48 h for MLB1.
7. Carefully remove the virus inoculum by tapping plates against 5–10 layers of paper towels, then remove paper towels into the autoclave bag. Alternatively, use a multichannel pipette to collect the virus-containing media into a plastic beaker containing 10% chloros to inactivate the virus in the safety cabinet. Follow local rules for the disposal of inactivated pathogens.
8. Fix the cells with 4% PFA in PBS (100 μL per well) for 20–30 min at room temperature. Disinfect the lid and borders of the 96-well plate with 4% PFA.
9. Before removing the plates from the safety cabinet, clean the outside of the plate with 1% Perasafe or 5% Chemgene.
10. Discard PFA after 30 min of fixing and wash the fixed cells with 1× PBS (100–200 μL per well). At this point, the plate can be stored at 4 °C or proceed directly for the immunostaining.
11. Permeabilize the fixed cells with 1% Triton X-100 (in 1× PBS) for 10 min on the rocker platform.
12. Wash the cells twice with 200 μL of 1× PBS.
13. Add 45 μL of diluted primary antibody per well to the cells and incubate on a rocker platform for 1–2 h at room temperature or 4 °C overnight. For MLB1/MLB2 virus capsid protein detection, use anti-MLB1 CP antibody (rabbit polyclonal, in-house, 1:300 is diluted in antibody solution). This antibody can be substituted with another custom-generated antibody against capsid-derived peptides.
14. Wash the cells three times with 200 μL of 1×PBS per well and incubate on the rocker for 5 min during each wash.
15. Add 45 μL of diluted secondary antibody (1:3,000) per well and incubate the plate on a shaker for 1–2 h at room temperature. Prepare anti-rabbit IRDye 800 secondary antibody in the antibody solution.
16. Discard the secondary antibody and wash cells three times with 200 μL of PBS per well; incubate on the rocker for 5 min during each wash.
17. Add 100 μL of PBS per well and clean the bottom of the plate with 70% ethanol.
18. Scan the plate on the LICOR imager using plate settings (+3 mm, medium, 84 μm). Each scan takes 25–30 min.
19. Export a 300 dpi image and quantify individual capsid-positive cells in the lowest dilution. Calculate the virus titer, taking the dilution factor into account. Using this method, titers of astroviruses are determined as infectious units per milliliter (IU/mL). An example of the MLB1 virus titration is shown in Figure 4.
Figure 4. Titration of MLB1 RG viruses. A. Huh7.5.1 cells are seeded on a 96-well plate, infected with 10-fold serial dilutions of MLB1 and fixed at 40–48 hpi, permeabilized, stained with anti-MLB1 capsid antibody, and imaged by Licor imager. B. Graph showing the titers of MLB1 viruses of 10 serial passages (P1 to P10). The MLB1 GNN mutant (GDD→GNN) harbors point mutations in the active site (GDD) of the viral RNA-dependent RNA polymerase (RdRp). Infectious units/mL = (No. of capsid-positive signals/0.01) × 1,000 μL/mL. Adapted from Ali et al. [1]. Data are mean ± SEM of 3 independent experiments.
Part II: Measuring MLB RG virus replication kinetics by western blotting
1. Seed Huh7.5.1 cells in the 6-multiwell plate (2.5 × 105/well) in 2 mL of complete media 24 h before infection.
Note: This should result in 60%–75% confluent cells.
2. Wash cells with 1 mL of DMEM-GHAA media.
3. Infect cells with MOI 1 of MLB1 and MLB2 viruses in a total volume of 250 μL of DMEM-GHAA media.
4. Incubate the cells for 1–2 h on a rocker at room temperature.
5. Add 1 mL of DMEM-GHAA-5% FBS media and carefully transfer the infected cells into the incubator (37 °C with 5% CO2).
6. Collect the infected cells at 24, 48, 72, and 96 h post infection. Since CPE can be present throughout the experiment, collect both media and cells by pipetting or scraping into a 1.5 mL Eppendorf tube, followed by centrifugation at 2,000× g for 5 min at 4 °C.
7. Discard the supernatants and resuspend the cell pellet in 80 μL of 50 mM Tris HCl pH 6.8, followed by the addition of 85 μL 2× LB-βME.
8. Immediately denature cell lysates by boiling at 95 °C for 5 min, followed by a spin for 1 min at 9,600× g.
9. Load 10 μL of cell lysates from each time point onto an 8%–12% SDS-PAGE gel and run the PAGE using standard running conditions (120 V for 1 h).
10. Transfer the resolved proteins to 0.2 μm nitrocellulose membranes using semi-dry transfer with a standard protocol (25 V, 1 A for 30 min).
11. Block the membrane with 4% milk in PBS for 1 h at room temperature.
12. Incubate the membrane with anti-MLB1 capsid antibody (rabbit polyclonal antibody, 1:3,000) and anti-tubulin (Abcam, ab6160, 1:1,000) for 1 h at room temperature or overnight at 4 °C on a rocker.
13. Wash the membrane three times with 0.1% PBST on a rocker for 5 min each time.
14. Add secondary antibodies, anti-rabbit (LI-COR IRDye 800, 1:3,000) and anti-mouse (LI-COR IRDye 680, 1:3,000), to the membrane and incubate for 1 h at room temperature or overnight at 4 °C on a rocker.
15. Wash the membrane three times with 0.1% PBST on a rocker for 5 min each time.
16. Scan immunoblot on a LI-COR ODYSSEY CLx imager and analyze using Image Studio version 5.2 (an example is shown in Figure 5).
Figure 5. Huh7.5.1 cells are infected with MLB1 and MLB2 viruses with MOI 1, viral capsid proteins were analyzed at the indicated time points using anti-CP MLB1 antibody (MLB1 CP antibody also detects MLB2 capsid protein). Anti-tubulin antibody is used as a loading control. This variation in tubulin levels can be explained by the combination of cell growth and virus-induced cell death and can be included in quantification if necessary.
Part III: Detection of virus infection by immunofluorescence
1. Day 1: Seed 5 × 104 Huh7.5.1 cells on IBIDI 8-well chambered slides (or grown on the sterile coverslips in a 12-multiwell plate at 1 × 105 cells per well).
2. Day 2: To infect the cells, first wash them with 1 mL of DMEM-GHAA media, and then infect with MOI 0.1 of MLB2 (or MLB1) virus in a total volume of 250 μL of DMEM-GHAA (keep one well of mock cells as a negative control).
3. Incubate cells for 1–2 h on a rocker at room temperature.
4. Remove virus inoculum and add 300 μL of fresh DMEM-GHAA-5% FBS media to each well. Then, carefully transfer infected cells to the incubator (37 °C with 5% CO2).
5. Fix the infected cells at 24, 48 and 72 h post infection. First, wash cells with 1× PBS, and then fix them with 10% formaldehyde in PHEM buffer (1:1) for 15 min. Alternatively, the cells can be fixed with 4% PFA in PBS.
6. Wash fixed cells three times with 1× PBS. At this stage, the fixed cells can be stored at 4 °C in PBS or directly proceed for immunofluorescence.
7. Permeabilize cells with 0.1% Triton X-100 for 10 min at room temperature.
8. Wash the cells three times with 1× PBS.
9. Block the cells with 2% goat serum in 1× PBS for 1 h at room temperature.
10. Add diluted primary antibodies (MLB1 anti-capsid (1:300) and anti-dsRNA IgG2a (Scicons J2, 1:250) to the cells and incubate them for 1 h at room temperature or overnight at 4 °C.
11. Wash the cells three times with 1× PBS to remove unbound antibodies.
12. Add diluted secondary antibodies (anti-rabbit Alexa 488- or anti-mouse 594-conjugated secondary antibody, 1:1,000) to the cells and incubate for 1 h at room temperature or overnight at 4 °C.
13. Wash the cells three times with 1× PBS to remove unbound antibodies.
14. Add 1:12,000 diluted Hoechst (in 1× PBS) to counterstain nuclei for 5 min at room temperature. Replace with 1× PBS.
15. Image MLB2 virus-infected cells using a Leica SP5 confocal microscope with a water-immersion 63Å objective or any similar confocal imaging platform.
An example of MLB2 virus infection/replication detection using this method is shown in Figure 6.
Figure 6. Representative confocal images of MLB2-infected Huh7.5.1 cells. MLB2 virus-infected cells are positive for capsid viral protein (in green), and virus replication is detected by using anti-dsRNA antibody (in red). Adapted from Ali et al. [2].
Part IV. Construction of MLB replicons and replicon assay
Replicon systems are considered valuable tools for the rapid measurement of RNA replication by inserting either fluorescent or luminescent reporter genes in place of the structural proteins coding sequence without altering the regulatory RNA elements essential for virus replication.
A. Engineering of MLB replicons
1. To generate MLB1 and MLB2 replicon systems, both the MLB1 (GenBank accession number ON398705) and MLB2 (GenBank accession number ON398706) RG clones are kept intact up to the end of ORFX, followed by a foot-and-mouth disease virus 2A sequence and a Renilla luciferase (Rluc) sequence with a stop codon, followed by the last 624 nucleotides of the virus genomes and a 35-nucleotide poly-A tail (Figure 7).
Figure 7. Schematic of the MLB replicons. The 2A-RLuc cassette is fused in the ORF2 followed by the stop codon and extended 3' UTR. SG indicates “subgenomic promoter.” Adapted from Ali et al. [1].
B. In vitro synthesis of replicon T7 RNA transcripts
1. Linearize replicon-containing plasmids (pMLB1R and pMLB2R, pMLB1R-GNN and pMLB2R-GNN) by digesting with the XhoI restriction enzyme.
2. Purify linearized plasmids using the PCR Clean-Up kit and elute in 20 μL of MilliQ water or elution buffer. Quantify concentrations of linearized plasmids using a Nanodrop.
3. Synthesize capped T7 RNA transcripts using T7 mMESSAGE mMACHINE Transcription kit (as shown in Table 7).
Table 7. In vitro synthesis of replicon T7 RNA transcripts
Components Volumes
1 Linear plasmids (100–200 ng) 1.5 μL
2 2× NTP/CAP 2.5 μL
3 10× transcription buffer 0.5 μL
4 T7 Enzyme mixture 0.5 μL
Total reaction volume 5.0 μL
4. Incubate the T7 RNA synthesis reactions at 37 °C for 60–90 min.
5. Add DNase I (2 U) for 20 min at room temperature to remove template plasmid DNA.
6. Check the T7-transcribed RNA(s) integrity by running on 1% agarose gel before or after DNase I digestion (Figure 8B).
7. Purify the T7-transcribed RNA(s) using an RNA Clean-Up and Concentrator kit and quantify concentrations using a Nanodrop, adjusting RNA concentration to 100–200 ng/μL.
C. Transfection of replicon T7-transcribed RNA(s)
1. Grow the Huh7.5.1 cells until they reach 70%–90% confluency (a T175 flask is sufficient for two 96-well plates).
2. Wash the cells with 10 mL of 1× PBS.
3. Trypsinize cells using 2–3 mL of 0.08% trypsin-EDTA at room temperature for 2–3 min.
4. Add 10 mL of DMEM-GH-2% FBS to neutralize trypsin activity and collect cells into 50 mL Falcon tubes.
5. Centrifuge the cells at 400× g for 5 min (with no brake, centrifugation takes more time to stop; so, in the meantime, start preparing transfection mixtures).
6. Discard the media and resuspend cells in 25 mL of DMEM-GH media.
7. Centrifuge the cells again at 400× g for 5 min (with no brake).
8. Discard the media and resuspend the cells in fresh DMEM-GH media to obtain a concentration of 1 × 106 cells/mL (at this point, cells are ready for transfection, so it is recommended to proceed immediately).
9. Prepare the transfection mixtures in the transfection media [serum-free OptiMEM containing RNaseOUT (40 units/mL)]. For mix 1, add (number of samples + 10%) × 0.5 μL of Lipofectamine 2000 and (number of samples + 10%) × 0.5 μL of OptiMEM in a 1.5 mL tube, mix well, and incubate for 5 min at room temperature. For mix 2, add 100 ng of T7-transcribed RNA(s) [90 ng of replicon T7-transcribed RNA(s) + 10 ng of T7-transcribed RNA(s), Renilla firefly internal control] in 10 μL of OptiMEM. For internal control, Firefly luciferase gene cloned under T7 promoter (Promega or similar) is used (cap-dependent translation should be consistent between replicates/samples).
10. After 5 min, combine mixes 1 and 2 and incubate them for 20 min at room temperature.
11. Add 100 μL of the prewashed cells (105 cells per well) to the transfection mixture and incubate at room temperature for 2–3 min.
12. Add FBS (5% final concentration) and then quickly transfer the cells into a flat-bottom 96-well plate.
13. Incubate transfected cells in a 37 °C incubator with 5% CO2.
14. Discard the media from cells at 4, 20, 24, and 30 h post-transfection and lyse the cells in the freshly prepared 1× passive lysis buffer (100 μL per well). To ensure complete cell lysis, freeze the plates at -70 °C for at least 30 min.
15. Thaw the plates at room temperature for 30 min and measure luciferase activity.
D. Measuring replicon activity by luciferase assay
1. To measure the luciferase reporter activity, take 20 μL of cell lysates and add 20 μL of Luciferase Assay Reagent II, followed by measurement of Firefly luciferase activity using GloMax® Navigator microplate luminometer.
2. Then, add 20 μL of freshly made 1× Stop & Glo reagent and incubate away from light for 10 min, followed by measurement of Renilla luciferase activity.
3. Calculate replicon activity as the ratio of Renilla luciferase (subgenomic reporter) to Firefly luciferase [co-transfected control T7-transcribed RNA(s)]. Subtract the background (mock-transfected cells) from each reading. The activity of the GNN mutant is used as a negative control, indicating no replication but active initial translation. The pMLB1R-GNN and pMLB2R-GNN (GDD→GNN) mutants carry point mutations in the active site (GDD) of the viral RNA-dependent RNA polymerase (RdRp), an enzyme critical for the replication of viral genomes.
An example of MLB1 and MLB2 replicon assay is shown in Figure 8.
Figure 8. Strategy to rapidly assess the replication dynamics of MLB astroviruses. A. Plasmids containing MLB replicon with Renilla luciferase and Firefly luciferase under the T7 promoter are linearized and used as a template for in vitro T7 RNA synthesis, followed by transfection and measurement of luciferase activity. B. The T7 RNA transcripts are checked on the 1% agarose gel. C. MLB1 and MLB2 replicon activities (replication dynamics) are measured in Huh7.5.1 cells. The replicon activity is normalized over firefly luciferase and shown as a relative replicon activity. Data are mean ± SD of 3 independent experiments. A RdRp knockout mutant (GDD→GNN) is used as replication-deficient control.
Data analysis
Data are shown as mean ± SEM (n = 3 independent experiments).
Data are shown as mean ± SD (n = 3 independent experiments).
Validation of protocol
The reproducibility of the protocol has been described in the following research articles:
• Ali et al. [1]. Attenuation hotspots in neurotropic human astroviruses. Plos Biology.
• Ali et al. [2] The astrovirus N-terminal nonstructural protein anchors replication complexes to the perinuclear ER membranes. Plos Pathogens.
General notes and troubleshooting
Troubleshooting
Ensure that RNA quality is high, as poor quality can negatively affect transfection/electroporation efficiency.
Confirm the sequence of reverse genetics and replicon plasmids before using them in the experiments.
Acknowledgments
This protocol was adapted from Ali et al. [1] and is partially based on recent work by Ali et al. [2]. This work was funded by a Sir Henry Dale Fellowship (220620/Z/20/Z) from the Wellcome Trust and the Royal Society to V.L. For the purpose of Open Access, the authors have applied a CC BY public copyright license to any Author Accepted Manuscript (AAM) version arising from this submission.
Competing interests
The authors have declared that no competing interests exist.
References
Ali, H., Lulla, A., Nicholson, A. S., Hankinson, J., Wignall-Fleming, E. B., O’Connor, R. L., Vu, D. L., Graham, S. C., Deane, J. E., Guix, S., et al. (2023). Attenuation hotspots in neurotropic human astroviruses. PLoS Biol. 21(7): e3001815.
Ali, H., Noyvert, D., Hankinson, J., Lindsey, G., Lulla, A. and Lulla, V. (2024). The astrovirus N-terminal nonstructural protein anchors replication complexes to the perinuclear ER membranes. PLoS Pathog. 20(7): e1011959.
Geigenmüller, U., Ginzton, N. H. and Matsui, S. M. (1997). Construction of a genome-length cDNA clone for human astrovirus serotype 1 and synthesis of infectious RNA transcripts. J Virol. 71(2): 1713–1717.
Article Information
Publication history
Received: Sep 11, 2024
Accepted: Dec 10, 2024
Available online: Jan 19, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
How to cite
Category
Microbiology > Microbe-host interactions > Virus
Microbiology
Molecular Biology > RNA > RNA synthesis
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5,202 | https://bio-protocol.org/en/bpdetail?id=5202&type=0 | # Bio-Protocol Content
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Combined FLIM, Confocal Microscopy, and STED Nanoscopy for Live-Cell Imaging
MB Magalie Bénard
CC Christophe Chamot
DS Damien Schapman
AL Alexis Lebon
LG Ludovic Galas
In Press, Available online: Jan 09, 2025
DOI: 10.21769/BioProtoc.5202 Views: 24
Reviewed by: Ivonne Sehring Anonymous reviewer(s)
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The authors used this protocol in Life Science Alliance Apr 2024
Abstract
Time-lapse fluorescence microscopy is a relevant technique to visualize biological events in living samples. Maintaining cell survival by limiting light-induced cellular stress is challenging and requires protocol development and image acquisition optimization. Here, we provide a guide by considering the quartet sample, probe, instrument, and image processing to obtain appropriate resolutions and information for live cell fluorescence imaging. The pleural mesothelial cell line H28, an adherent cell line that is easy to seed, was used to develop innovative advanced light microscopy strategies. The chosen red and near-infrared probes, capable of passively penetrating through the cell plasma membrane, are particularly suitable because their stimulation from 600 to 800 nm induces less cytotoxicity. The labeling protocol describes the concentration, time, and incubation conditions of the probes and associated adjustments for multi-labeling. To limit phototoxicity, acquisition parameters in advanced confocal laser scanning microscopy with a white laser are determined. Light power must be adjusted and minimized at red wavelengths for reduced irradiance (including a 775 nm depletion laser for STED nanoscopy), in simultaneous mode with hybrid detectors and combined with the fast FLIM module. These excellent conditions allow us to follow cellular and intracellular dynamics for a few minutes to several hours while maintaining good spatial and temporal resolutions. Lifetime analysis in lifetime imaging (modification of the lifetime depending on environmental conditions), lifetime dye unmixing (separation with respect to the lifetime value for the spectrally closed dye), and lifetime denoising (improvement of image quality) provide flexibility for multiplexing experiments.
Key features
• Cell preservation after labeling with less cytotoxic red, near-infrared dye viable probes.
• Determination of lower but efficient probe concentration; adjust good balance between probes concentration and incubation time to achieve multi-labeling.
• Long time-lapse acquisition in advanced confocal microscopy with sensitive new-generation detectors.
• Confocal image combined with fast FLIM for multi-labeling with spectrally closed dyes, unmixed from lifetime values.
• Confocal-STED image acquisition combined with fast FLIM to improve signal-to-noise ratio.
Keywords: Live-cell imaging Sample preservation Fluorescence lifetime Confocal Fast FLIM STED
Graphical overview
Background
Reducing cell exposure to study cellular dynamics is essential for cell biology research. As previously described [1], the quartet sample, probe, instrument, and image processing represent essential steps to conduct fluorescence microscopy experiments.
To preserve the viability of cells during imaging, the photophysical properties of dyes must be considered. In particular, their photostability under light excitation and brightness, with high quantic yield to reduce the light power to a minimum. The incorporation of the probe must be specific, efficient, and non-toxic inside the cell. The protocol must be optimized to determine the minimum effective concentration to strongly label the cell elements by limiting background noise. For multi-labeling, the concentration and incubation time balance must be optimized [2]. The choice of red/near-infrared dyes is compatible with living cells (red excitation light is less irradiating than lower wavelength illumination). The dye must be depletable with a 775 nm laser to perform STimulated-Emission-Depletion (STED) acquisition in nanoscopy.
To image cellular compartments, red fluorescent dyes linked to ubiquitous markers (e.g., wheat germ agglutinin Alexa Fluor 633 conjugate, AF633-WGA for lectin, Nile red for lipid, MitoTracker Red CMXRos, MTR and LBL-Dye M715 for mitochondria, SPY620-DNA for nucleus, SPY650-FastAct and SiR700-Act-Verapamil for actin cytoskeleton) were used for single and multiple labeling experiments. Their depletion capacity at 775 nm was verified and the lifetime characteristics (value and mono- or multi-components after exponential curve fitting) were determined [2,3].
To perform time-lapse multiplex acquisition by limiting light exposure, the dyes, all of which have a near-red wavelength, are stimulated and detected in one detector. Their specific lifetime values allow their separation in several fluorescence channels. The tunable white light laser (WLL) provides high flexibility in wavelength adjustment; the pulsed 775 nm laser depletes the red/near-infrared dyes in STED nanoscopy and induces less irradiance of living cells compared to other continuous depletion lasers [2,4]. The new generation of high-speed photon counting detectors, hybrid detectors (HyD), brought striking opportunities for high signal-to-noise ratios and fast detection of fluorescence signals, particularly true for HyD-X within the red/near-infrared range [2]. Biologist-friendly fluorescence lifetime imaging microscopy (FLIM), called fast FLIM [5–7], combined with confocal or confocal STED acquisition image, offers a flexible method suitable for cellular preservation and provides the lifetime parameter to xyzt dimensions while maintaining axial and temporal resolution.
Lifetime dye unmixing is an alternative to the limited fluorescence spectral separation. Simultaneous acquisition (one step), which is fast and limits cell light exposure compared to sequential acquisition, followed by lifetime dye unmixing allows (i) signal denoising by lifetime value selection to eliminate background or nonspecific signal, and (ii) dye fluorescence selection on the phasor, representing the photon lifetime distribution [8]. This method of multiplexing living cells can be achieved by activating one and eventually several detectors [9], which is more limited in STED nanoscopy. As the lifetime value can be influenced by factors such as viscosity, temperature, and pH [10–11], the lifetime imaging is informative about the cellular environment, revealing multi-components after fitting of the fluorescence decay curve; the resolution gain in STED microscopy can be improved in lifetime denoising modality by removing uncorrelated STED process photons and smoothing the signal. This protocol is applicable to all types of adherent cells and viable red/near-infrared dye probes.
Materials and reagents
Biological materials
1. Human H28 cell line (ATCC, NCI-H28, catalog number: CRL-5820) from the lungs of a 48-year-old white male with stage 4 mesothelioma.
Reagents
1. Roswell Park Memorial Institute (RPMI) 1640 cell culture medium (Thermo Fisher Scientific, Gibco, catalog number: 31870-025)
2. L-Glutamine 200 mM, 100× (Thermo Fisher Scientific, Gibco, catalog number: 25030-024)
3. Fetal bovine serum (FBS) (Life Technologies, Gibco, catalog number: 10270-106), heat-inactivated at 56 °C for 2 h
4. Antibiotic/antimycotic solution 100× with 10,000 U/mL penicillin G, 10,000 μg/mL streptomycin, and 25 μg/mL amphotericin B (HyClone, catalog number: SV30079.01)
5. Trypsin-EDTA with phenol red (0.25%) (Fisher Scientific, catalog number: 11560626)
6. Phosphate buffered saline (PBS), pH 7.4, 10× solution (Thermo Fisher Scientific, Gibco, catalog number: 70011-044)
7. Distilled water (Thermo Fisher Scientific, Invitrogen, catalog number: 10977-035)
8. Dimethyl sulfoxide (DMSO) (Corning, catalog number: 25-950-CQC)
9. Methanol anhydrous (Carlo Erba, catalog number: P09310D16)
10. Wheat germ agglutinin Alexa Fluor 633 conjugate (AF633-WGA) (Thermo Fisher Scientific, Invitrogen, catalog number: W21404)
11. Nile Red (Merck, Sigma-Aldrich, catalog number: N3013)
12. MitoTracker Red CMXRos (MTR) (Thermo Fisher Scientific, Life Technologies, catalog number: M7512)
13. LBL-DyeM 715 (Proimaging, catalog number: BM298)
14. SPY650-FastAct (Spirochrome, catalog number: SC505)
15. SPY620-DNA (Spirochrome, catalog number: SC404)
16. SiR700-Act and Verapamil kit (Spirochrome, catalog number: SC013)
Laboratory supplies
1. Cell culture flask 25 cm2, T-25 (TPP, catalog number: 90026)
2. 10 mL serological pipette (Merck, Costar, catalog number: CLS4488)
3. 5 mL serological pipette (Merck, Costar, catalog number: CLS4487)
4. 2 mL serological pipette (Merck, Costar, catalog number: CLS4486)
5. 15 mL Falcon tube (Falcon, catalog number: 352096)
6. 50 mL Falcon tube (Falcon, catalog number: 352070)
7. 1.5 mL microcentrifuge tubes (Eppendorf, catalog number: 022431081)
8. Glass-bottom microwell dishes, 35 mm Petri dish, 20 mm microwell, No. 1.5 coverglass 0.16–0.19 mm (MatTek Corporation, catalog number: P35G-1.5-20-C)
9. Neubauer chamber (EMS, catalog number: 68052)
10. Dropping bottles (Avantor, catalog number: HECH41314050)
Equipment
1. Class II biosafety cabinet (Thermo Scientific, catalog number: 51028226, model: MSC-ADVANTAGE)
2. CO2 incubator (Thermo Scientific, catalog number: 51032874, model: Heracell 150)
3. Heating oven (Thermo Scientific, catalog number: 51028112, model: Heratherm)
4. Safe liquid aspiration system Vacusafe (Avantor, Integra Biosciences, catalog number: 391-2094, model: Vacusafe Comfort plus)
5. Ultrasonicator (Ultrasonik Ney, model: 19H)
6. Pipette controller (Thermo Fisher Scientific, Integra Biosciences, catalog number: 155017, model: Pipetboy acu 2)
7. Mechanical pipette 0.1–2.5 μL (Eppendorf, catalog number: 3123000012, model: Research Plus)
8. Mechanical pipette 0.5–10 μL (Eppendorf, catalog number: 3123000020, model: Research Plus)
9. Mechanical pipette 20–200 μL (Eppendorf, catalog number: 3123000055, model: Research Plus)
10. Mechanical pipette 100–1000 μL (Eppendorf, catalog number: 3123000063, model: Research Plus)
11. Centrifuge for 15 mL and 50 mL tubes (Thermo Fisher Scientific, Eppendorf, catalog number: 5804000528, model: 5804R)
12. Microcentrifuge for tubes 1.5 mL (Thermo Fisher Scientific, Fisherbrand, catalog number: 75002460, model: Micro 17)
13. Power meter Argo-Power Slide (Argolight, model: Argo-POWER Slide V2)
14. Culture cell observation microscope (Leica Microsystems, model: DMIL LED)
15. Inverted stand confocal laser scanning microscope (Leica Microsystems, model: STELLARIS 8, FALCON STED 3 X WLL) equipped with a white light laser (WLL, 440–790 nm), 775 nm depletion laser for STED in living cells, four hybrid detectors (HyD type S, X, and R), an 86× objective (NA = 1.20, water immersion, WD = 300 µm), a fully fast integrated FLIM module FAst Lifetime CONtrast (FALCON, Leica Microsystems), and a full bold line Okolab chamber (Ottaviano, Italy) to keep the temperature at 37 °C and mix the CO2 at 5%
Software and datasets
1. Leica Application Suite X (LAS X, Leica Microsystems, version 4.5.0)
2. Leica Application Suite X FLIM/FCS (LAS X, Leica Microsystems, version 4.5.0)
3. Leica Application Suite X 3D viewer (Leica Microsystems, version 4.5.0)
4. Fiji/ImageJ (open source, National Institutes of Health, [12])
Procedure
A. Cell seeding
1. Culture H28 cells in complete RPMI cell culture medium [RPMI cell culture medium supplemented with 10% FBS, 1% glutamine (2 mM), and 1% antibiotic/antimycotic solution] at 37 °C in a humidified atmosphere with 5% CO2. Grow cells in a T-25 flask until confluence before experiments (maximum 90% confluence).
2. Remove the media by aspiration with the safe liquid aspiration system from the cell culture flask and briefly wash the cells with prewarmed PBS (5 mL, 37 °C).
3. Remove the PBS by aspiration with the safe liquid aspiration system and apply 2 mL of prewarmed trypsin solution (37 °C).
4. Incubate the cells at 37 °C for 2 min in the CO2 incubator.
5. Add 5 mL of prewarmed complete RPMI cell culture medium and dissociate the cells by pipetting.
6. Transfer the dissociated cells to a 50 mL Falcon tube and centrifuge at 1,000× g for 5 min at room temperature.
7. Remove the supernatant by using the safe liquid aspiration system at low speed and resuspend the cells in 5–10 mL of complete RPMI cell culture medium.
8. Determine the number of cells per milliliter using a Neubauer chamber.
9. Plate 400,000 cells per cm2 in 35 mm glass-bottom microwell dishes in 2 mL of complete RPMI cell culture medium and allow the cells to incubate for 1–2 days before staining and imaging.
B. Probe preparations and cell staining
1. Probe preparation for storage (1,000× stock solutions)
a. Dissolve wheat germ agglutinin Alexa Fluor 633 conjugate (AF633-WGA) powder in DMSO to obtain a 1 mg/mL stock solution. Aliquot 2 μL into 1.5 mL tubes and store at −20 °C.
b. Resuspend Nile Red powder in pure methanol to obtain a 1 mg/mL stock solution. If aggregates are visible in the solution, sonicate for 2–5 min until the preparation is completely dissolved. Store the stock solution tube at 4 °C.
c. Dissolve MitoTracker Red CMXRos (MTR) powder in DMSO to obtain a 1 mM stock solution. Aliquot 1 μL into 1.5 mL tubes and store at −20 °C
d. Dissolve LBL-Dye M715 powder in DMSO to obtain a 2 mg/mL stock solution. Aliquot 2 μL into 1.5 mL tubes and store at −20 °C. As the fluorescence intensity of LBL-Dye M715 is strong, it may be necessary to dilute this solution up to 4-fold.
e. Prepare Spirochrome probes as directed by the supplier by dissolving SPY650-FastAct and SPY620-DNA in 50 µL of DMSO to obtain a 1 mM stock solution stored at −20 °C. For SiR700-Act, supplement the 35 nmol with 100 µL of DMSO to obtain a 1 mM stock solution. One tube of 10 mM Verapamil (1,000× stock solution) is included in the SiR700-Act kit. Do not divide the solution into small aliquots, as indicated in the supplier’s datasheet.
2. Probe preparation with extemporaneous dilution for cell staining
a. One to two days after seeding adherent cells onto 35 mm glass-bottom microwell dishes, cell density (40%–60% cell confluence/600–800,000 cells per cm2), physiology state (shape and proliferation of the cells), and adherence are optimal for imaging. Freshly prepare probe(s) solutions as follows:
b. For single labeling of cells with AF633-WGA, MTR, and LBL-Dye M715, freshly dilute the probe stored in a frozen stock tube at −20 °C in 1 mL of prewarmed (37 °C) complete RPMI cell culture medium to obtain a 2× solution.
c. For single labeling of cells with Nile Red staining, transfer 2 μL of probe stored at 4 °C in 1 mL of prewarmed (37 °C) complete RPMI cell culture medium to obtain a 2× solution. For long-time acquisition (more than 30 min), the Nile Red solution can be diluted 2 or 3-fold to limit dye accumulation.
d. For single labeling with SPY 650-FastAct or SPY 620-DNA, transfer 1–2 μL of probe in 1 mL of prewarmed (37 °C) complete RPMI cell culture medium to obtain a 2× solution.
e. For single labeling of cells with SiR700-Act, transfer 1–2 μL of probe supplemented with 2 μL of Verapamil solution, to 1 mL of prewarmed (37 °C) complete RPMI cell culture medium to obtain a 2× solution.
f. For multi-labeling of cells, the probes can be mixed in the same final tube containing 1 mL of prewarmed (37 °C) complete RPMI cell culture medium to obtain a 2× solution for both or all three probes. Probes are added directly into the cell culture medium depending on the minimum incorporation time (see step B3).
3. Cell staining for image acquisition
a. Remove the medium from adherent cells onto 35 mm glass bottom microwell dishes.
b. Gently apply 1 mL of prewarmed (37 °C) complete RPMI cell culture medium to the edge of the dishes.
c. Transfer 1 mL of the diluted probe(s) 2× solution onto the cells, very gently, drop by drop.
d. Incubate H28 cells with the probe(s) inside the incubator (37 °C/5% CO2) for:
- 10 min minimum with Nile Red probe(s),
- 5–10 min with AF633-WGA to observe the incorporation of the dye at the plasma cell membrane level; 15 min minimum to observe the internalization of WGA for a few hours afterward in the cell cytoplasm,
- 30 min minimum for MTR and LBL-Dye M715,
- 1 h minimum for SPY620-DNA, SPY650-FastAct, and SiR700-Act.
e. In case of complex multi-staining such as MTR combined with SiR700-Act or SPY620-DNA, or SiR700-Actin with LBL-Dye M715, a first incubation is performed with probes requiring long-time incubation (SiR700-Act, SPY620-DNA, SPY650-FastAct) for 30 min followed by another incubation of 30 min after adding directly the stock probe (1 μL of MTR or 2 μL of LBL-Dye M715, as indicated in steps B1c and B1e) to the cell culture dish and mixing gently by pipetting.
f. Observe the cells directly under the microscope without removing the medium, unless there is a high level of nonspecific fluorescence signal (see Troubleshooting, Problem 4).
C. Microscope setup for live cell imaging
1. Turn on the instrument (microscope controller, laser power, scanning laser head, FLIM module) and the computer.
2. After 10 min, open the LAS-X software.
3. If STED setup is to be used, select the option.
4. One or two hours before acquisition, warm up the chamber installed around the inverted microscope stand to the temperature of 37 °C.
5. Turn on the laser boxes and preheat the laser source to 85%–100% for the WLL laser (excitation laser).
Note: As previously described [2, Figure S1], laser powers of 594, 638, and 685 nm at 100% AOTF are measured at 523, 386, and 382 µW ±4.5, respectively, without the objective in the light path through the optical power meter integrated in Argo-Power slide.
6. If the STED laser is to be used, turn on the 775 nm STED depletion laser at 100% (557 mW measured without the objective in the light path for 100% power setting).
7. Place the 86× water immersion objective suitable for live cell observation and apply a drop of distilled water to the surface of the objective lens by using a dropping bottle of distilled water.
8. Place the glass bottom microwell dish on the microscope stage and lift the objective until contacting with the water drop.
9. Place the cover in a humidified atmosphere with 5% CO2.
10. Optimize the acquisition parameters: conventional scanner at 400 Hz, format 1024 × 1024 pixels, and pixel dwell time 1.4 μs (see Figure 1A). To improve the acquisition speed, the scanning frequency can be increased to 600 Hz (pixel dwell time decreased to 875 ns) and in bidirectional mode (pixel dwell time: 600 ns). The low pixel dwell time can be compensated by line repetition to accumulate photons per pixel. The format can be reduced to 512 × 512 pixels if the biological element to observe does not require a small pixel size (pixel dwell time increased to 1.2 μs with format 512 × 512, scan rate 600 Hz, bidirectional scanning mode). An optimal standard Airy 1 pinhole is used by default.
11. Select the xyzt acquisition mode (see Figure 1B).
12. Find focus and select an appropriate field of view with the microscope eyepiece.
Figure 1. Screenshot of LAS X software for image acquisition of three dyes simultaneously. In this example, three probes (SPY620-DNA, SiR700-Act, and LBL-Dye M715), indicated by * in Table 1, are acquired in one single detector after 618 nm laser excitation (5% WLL laser power). The confocal image, on the right side, shows one color image for the three dyes.
D. Simultaneous confocal/confocal-STED image acquisition with fast FLIM module activation
1. Determine the excitation wavelength [power minimum between 2% and 6% of acousto-optic tunable filter (AOTF)] as mentioned in Table 1.
2. Activate the adapted detector for red dye(s), the HyD-X, in photon counting mode, and determine the emission band (see Figure 1C).
Note: Other detectors can be activated, in simultaneous mode, for multi-channel acquisition.
Table 1. Excitation and emission wavelengths for dye(s) detection in unique HyD-X detector.
Probe (dye) Exc (nm) Em (nm) in HyD-X detector
AF633-WGA 633 640–670
Nile Red 600 615–670
MTR 590 605–630
LBL-Dye M715 690 705–740
SPY650-FastAct 650 660–715
SPY620-DNA 618 625–640
SiR700-Act 698 710–740
SPY650-FastAct + LBL-Dye M715 650 660–740
SPY620-DNA + SiR700-Act + LBL-Dye M715 (*) 618 625–740
3. In STED mode, activate the 775 nm at low power (2%–20 %) for depletion effect (Figure 1, yellow arrow).
Note: As the lifetime characteristics of dyes are deeply modified by the depletion laser [13], it is recommended to use a separate detector for each dye in multi-labeling experiments.
4. Adapt the zoom factor and pixel size in coherence with the sample.
Note: In STED mode, since spatial resolution is often the priority, it is necessary to zoom in until reaching an optimal pixel size of 20 nm. In this case, a strong accumulation of photons (by reasonably increasing the excitation laser power and preferably by adding scanning line accumulation) is essential to compensate for the reduced pixel size and to have sufficient signal to use the FLIM module.
5. Determine the Z up and down in case of z-stack acquisition.
6. Determine the time interval and total duration for time-lapse.
7. Activate the fast-integrated FLIM module (see the button in Figure 1D), the so-called FAst Lifetime CONtrast (FALCON), with 1–3 line repetitions to reach a minimum of 50,000 photons per image.
Note: If the spatial resolution is not a crucial point, enlarging the pinhole can be an alternative to line repetitions.
8. Acquire the confocal or confocal-STED image(s) (Figure 1E) by clicking Start.
9. A FLIM image appears in the second window in the LAS X FLIM/FCS software.
E. Visualization of fast FLIM images
1. Data from the LAS X FLIM/FCS software are divided into three main parts:
a. “FLIM” for lifetime imaging with FLIM fitting curve measurement or fast FLIM images (Figure 2A).
b. “Phasor” FLIM image for lifetime imaging and phasor separation (lifetime dye unmixing; Figure 2B).
Note: The lifetime value is modified by the STED depletion laser and can be difficult to unmix.
c. “τ-STED” for lifetime denoising of STED image only (Figure 2C).
2. The lifetime image can be directly exported by clicking on the Save image button (Figure 2, yellow arrow).
3. Lifetime dye unmixing images are produced after Phasor separation: select the draw cursor corresponding to the lifetime values of each dye (Figure 3A). Click on Separate (Figure 3B) and visualize the lifetime separation in fluorescence channels (Figure 3C). Save the image by clicking the corresponding button (Figure 3, yellow arrow).
Note 1: It is necessary to control/measure the value for each dye separately to determine their respective positions on the Phasor plot representation.
Note 2: A minimum lifetime difference of 0.4 ns seems to be necessary to spatially separate the dyes.
4. The τ-STED option (Figure 2C) is available for images acquired after STED laser activation. Choose the τ-strength and denoise level (Figure 4B) depending on dye signal emission (quality of biologic element observed; level of nonspecific signal and background). Lifetime denoising images can be combined with lifetime images acquired from two separate detectors (one with lifetime information and the other with lifetime denoising improvement).
5. All these acquisitions and data FLIM analysis can be performed in z-stack and time-lapse (xyzt), as seen in Figure 4, with z and t lines (yellow arrows). In this example, the simultaneous 2-channels acquisition is performed through time-lapse containing 60 time points, one image every 10 s for 10 min, in a 4-z plan. Tracking and fluorescence intensity can be measured using the Manual Tracking plugin of Fiji software (see Figure 5).
6. All images are exported and saved through the Save image button in .lif format (Leica) and can be opened and analyzed with Fiji software (see Figure 6).
Figure 2. Screenshot of LAS X FLIM/FCS window for FLIM data. The three main parts for data FLIM analysis are FLIM (A), Phasor (B), and τ-STED (C) analysis. Note that the τ-STED button is available only for confocal-STED images. In this Phasor window, the lifetime image is visualized in D, with a possibility to modify the scaling in intensity (black and white bar, E) and the scaling of lifetime values (rainbow bar, F). This image is obtained during confocal acquisition (Figure 1, with detector of SPY620-DNA, SiR700-Act, and LBL-Dye M715 in Hyd-X detector). Lifetime values of each pixel are distributed in Phasor plot (G).
Figure 3. Lifetime dye unmixing of three dyes from the same confocal image. From the confocal image (Figure 1), the lifetime separation through the Phasor (Figure 3A) produced three channels (Figure 3C) corresponding to the separation of three lifetime value zones (A, left white circle, Tau 3.2 ns, SPY620-DNA; A, middle white circle, Tau 1.7 ns, SiR700-Act; A, right white circle, Tau 1.1 ns, LBL-Dye M715).
Figure 4. Lifetime imaging and lifetime denoising information after sequential data treatment of two dyes signal in two detectors. In this example, STED acquisition is realized from cells double-labeled with MTR (channel 1, Exc 590, 1% WLL laser power, HyD-S 605–630 nm) and SiR700-Act (channel 2, Exc 698 nm, 6% WLL laser power, HyD-X 710–740 nm). Window A shows the lifetime image of MTR and SiR700-Act; the FLIM analysis can be realized in τ-STED mode to improve image quality (lifetime denoising, B). The Denoise parameter at “50” is sufficient for MTR; the background is higher for SiR700-Act, and the value can be increased to 70 to improve the resulting image after lifetime denoising. The combination of these two distinct FLIM analyses is possible (MTR with its informative lifetime image; SiR700-Act with the thin structure of actin improved by FLIM denoising).
Figure 5. Tracking analysis illustration with ImageJ Macro. Time-lapse of mitochondrial dynamics in H28 cells labeled with LBL-Dye M715 and STED acquisition followed by lifetime denoising processing (Exc 690 nm, 3% WLL laser power, HyD-X 705–740 nm, 2% STED laser 775 nm). Tracking and fluorescence intensity are measured using the Manual Tracking plugin of Fiji software [12]. Trajectory (A) is drawn in Fire LUT in the final image (t35 min) and the corresponding graph representing the distance traveled by the element from the origin point (t0, *) in function of time (35 min of acquisition) is shown in B. The white arrow represents the direction of mitochondrial movement.
Figure 6. Fiji/ImageJ to import and analyze dye unmixing. From the confocal image (Figure 1), the lifetime dye unmixing of three dyes separated in three channels, Fiji/ImageJ software allows the exportation of the image in .tif format, histogram scaling, and LUT attribution to obtain a definitive image with a scale bar.
Data analysis
1. Fiji/ImageJ to import and analyze lifetime images:
a. Open the image stack with Fiji/ImageJ software (including the plugin bioformats).
b. Change the order of the look-up table (LUT) from red-green-blue (RGB) in .lif format to blue-green-red to restore the correct order lifetime LUT.
c. Save as RGB image in .tiff
2. Fiji/ImageJ to import and analyze dye unmixing:
a. Open the image stack with Fiji/ImageJ software (including the plugin bioformats).
b. Image > Color > Split Channels to separate images from stack.
c. Image > Adjust > Brightness/Contrast to modify the fluorescence histogram scaling. This step is important to restore the signal balance between the fluorescence channels.
d. Image > Lookup Tables to choose the adapted color combination to reveal dye information.
e. Image > Color > Merge Channels to produce an overlay image.
f. Image > Type > RGB Color to fuse the three fluorescence channels in one resulting image.
g. Analyse > Tools > Scale bar to obtain the definitive image to save in .tiff format.
h. File > Save as > Tiff.
3. ImageJ Macro, Tracking analysis:
a. For image processing and analysis, use Fiji or ImageJ software. In the latter case, please ensure that you have the plugins bioformats and manual tracking installed.
b. Open your timelapse image and check in image properties (Image > Properties) if the frame interval is correct.
c. Launch the manual tracking plugin (https://imagej.net/ij/plugins/track/Manual%20Tracking%20plugin.pdf). In the very bottom part of the tracking window, tick the show parameters case. Fill time interval and XY calibration with the image properties.
d. Add your first track. Once you have clicked on the add track button, you can go back to your timelapse image. Each time you click on it, you will go to the next slice, and the results table will be updated. Finish your track with the end track button.
e. Now you can generate a trajectory with the progressive lines button.
f. To represent time as color on a projection, you can manipulate this progressive line stack. In RGB images, first convert it to an 8-bit image in Image > Type > 8-bit.
g. All the points of the line should be at 85 in intensity. Each point should have its own value. To do so, we wrote a macro sequence:
run("32-bit");
run("Divide...", "value=85 stack"); //process the whole stack
for (i = 1; i <= nSlices; i++) {
setSlice(i);
run("Multiply...", "value=i slice");
}
h. The time series can now be projected in Progressive Lines with Image > Stacks > Z projection.
i. A fire LUT can be applied in Image > Look-up Tables > Fire. Adjust the brightness and contrast to fill the whole dynamics of the line with the range of the look-up table.
j. Finally, to paste the projection onto the appropriate image, change the type of both progressive line projection and destination image to RGB (Image > Type > RGB image).
k. Edit > Paste Control, choose Transparent-zero. Copy the progressive lines projection and paste it on the destination image.
Validation of protocol
This protocol or parts of it has been used and validated in the following research articles:
Bénard et al. [3]. Combining sophisticated fast FLIM, confocal microscopy, and STED nanoscopy for live-cell imaging of tunneling nanotubes. Life Sci Alliance (Figures 2–7 for protocol and Figures S3–5).
Bénard at al. [2]. Optimization of Advanced Live-Cell Imaging through Red/Near-Infrared Dye Labeling and Fluorescence Lifetime-Based Strategies. Int J Mol Sci (Figures 4–7 for protocol and Figures S1–S6 for quality assessment).
General notes and troubleshooting
General notes
This protocol was developed for the human H28 cell line and can be adapted for any adherent cell type. All probes and dyes, after identification of non-toxicity in living cells during time-lapse, verification of sufficient signal-to-noise ratio, and measurement of the lifetime value can be tested by this method. Spirochrome probes are known to be suitable for fluorescence microscopy of live cells and are tested on Hela and COS-7 cells [14]. However, some reagents can interfere with cellular physiology, including SPY-DNA, which binds DNA, and Ca2+ channel-inhibitor verapamil, which may affect cellular excitability and signaling. This protocol with sample preparation on adherent cells is compatible with any microscopy devices able to maintain cells alive and equipped with equivalent confocal/STED/FLIM systems.
Troubleshooting
Problem 1: Cell death.
Possible causes: Cell death can occur before the experiment (culture problems), during cell labeling (probe toxicity problems), or during microscope observation (light toxicity problems).
Solutions: Check the cell culture environment (medium, incubator, etc.). If the probe is new, check the viability (with NucGreen, death marker Exc488 nm, for example) and perform time and dose-response curves; reduce the dye concentration if the signal remains good. If cell death occurs during observation, check the exposure laser power; leave the labeled cells in the incubator in parallel with the observation to find out if the acquisition parameters are the cause.
Problem 2: The fluorescence signal is weak.
Possible causes: Check your system (laser power, detector, etc.); check your cells (a change in cell density or metabolic state can decrease the efficiency of probe incorporation); check your probe (some dyes may not be stored for a long time).
Solutions: Increase the probe concentration and/or incubation time if the cell density is higher (conditions to be adapted for other cell types); buy a new probe. Adding Verapamil, a broad-spectrum efflux pump inhibitor, or pluronic acid, known to dissolve dye aggregates, may help the probe to incorporate into the cell and increase the bioavailability of the dye, respectively. Use only freshly prepared staining solutions.
Problem 3: Unbalanced multiple labeling.
Possible causes: Dye to be renewed; cell no longer incorporates probes in the same way due to metabolic disturbance.
Solutions: Delicate balance. Change incubation time and/or concentration of low-fluorescence dyes. Adapt the excitation wavelength and emission band to find a better balance for emission signals.
Problem 4: High background noise.
Possible causes: Concentration of probe too high or emission signal too weak (boost the signal and revelation of the background noise); medium with phenol red as pH indicator (especially true with DMEM, which can produce autofluorescence).
Solutions: Adapt the concentration of the probe. Change the culture medium to remove the excess probe; use a culture medium without phenol red.
Acknowledgments
This work was supported by the University of Rouen Normandy, Inserm, IRIB, Région Normandie (RIN plate-forme “7D microscopy”, RIN émergent COMVOI), the European Regional Development Fund (ERDF “7D Microscopy”), the GIS IBiSA, and the national infrastructure France-BioImaging supported by the French National Research Agency (ANR-10-INBS-04).
Competing interests
The authors declare no competing interest.
References
Galas, L., Gallavardin, T., Bénard, M., Lehner, A., Schapman, D., Lebon, A., Komuro, H., Lerouge, P., Leleu, S., Franck, X., et al. (2018). “Probe, Sample, and Instrument (PSI)”: The Hat-Trick for Fluorescence Live Cell Imaging. Chemosensors. 6(3): 40. https://doi.org/10.3390/chemosensors6030040
Bénard, M., Schapman, D., Chamot, C., Dubois, F., Levallet, G., Komuro, H. and Galas, L. (2021). Optimization of Advanced Live-Cell Imaging through Red/Near-Infrared Dye Labeling and Fluorescence Lifetime-Based Strategies. Int J Mol Sci. 22(20): 11092. https://doi.org/10.3390/ijms222011092
Bénard, M., Chamot, C., Schapman, D., Debonne, A., Lebon, A., Dubois, F., Levallet, G., Komuro, H. and Galas, L. (2024). Combining sophisticated fast FLIM, confocal microscopy, and STED nanoscopy for live-cell imaging of tunneling nanotubes. Life Sci Alliance. 7(7): e202302398. https://doi.org/10.26508/lsa.202302398
Leutenegger, M., Eggeling, C. and Hell, S. W. (2010). Analytical description of STED microscopy performance. Opt Express. 18(25): 26417. https://doi.org/10.1364/oe.18.026417
Alvarez, L. A., Widzgowski, B., Ossato, G., van den Broek, B., Jalink, K., Kuschel, L., Roberti, M. J. and Hecht, F. (2019) Application Note : SP8 FALCON : a novel concept in fluorescence lifetime imaging enabling video-rate confocal FLIM. Nat Methods. https://www.nature.com/articles/d42473-019-00261-x
Bitton, A., Sambrano, J., Valentino, S. and Houston, J. P. (2021). A Review of New High-Throughput Methods Designed for Fluorescence Lifetime Sensing From Cells and Tissues. Front Phys. 9: e648553. https://doi.org/10.3389/fphy.2021.648553
Gonzalez Pisfil, M., Nadelson, I., Bergner, B., Rottmeier, S., Thomae, A. W. and Dietzel, S. (2022). Stimulated emission depletion microscopy with a single depletion laser using five fluorochromes and fluorescence lifetime phasor separation. Sci Rep. 12(1): e1038/s41598–022–17825–5. https://doi.org/10.1038/s41598-022-17825-5
Digman, M. and Gratton, E. (2012) Complex fluorescence decay profiles: discrete models, stretched exponential, analytical models. In Fluorescence Lifetime Spectroscopy and Imaging: Principles and Applications in Biomedical Diagnostics. https://escholarship.org/uc/item/5g279175
Frei, M. S., Tarnawski, M., Roberti, M. J., Koch, B., Hiblot, J. and Johnsson, K. (2021). Engineered HaloTag variants for fluorescence lifetime multiplexing. Nat Methods. 19(1): 65–70. https://doi.org/10.1038/s41592-021-01341-x
Berezin, M. Y. and Achilefu, S. (2010). Fluorescence Lifetime Measurements and Biological Imaging. Chem Rev. 110(5): 2641–2684. https://doi.org/10.1021/cr900343z
Datta, R., Heaster, T. M., Sharick, J. T., Gillette, A. A. and Skala, M. C. (2020). Fluorescence lifetime imaging microscopy: fundamentals and advances in instrumentation, analysis, and applications. J Biomed Opt. 25(7): 1. https://doi.org/10.1117/1.jbo.25.7.071203
Schindelin, J., Arganda-Carreras, I., Frise, E., Kaynig, V., Longair, M., Pietzsch, T., Preibisch, S., Rueden, C., Saalfeld, S., Schmid, B., et al. (2012). Fiji: an open-source platform for biological-image analysis. Nat Methods. 9(7): 676–682. https://doi.org/10.1038/nmeth.2019
Tortarolo, G., Sun, Y., Teng, K. W., Ishitsuka, Y., Lanzanó, L., Selvin, P. R., Barbieri, B., Diaspro, A. and Vicidomini, G. (2019). Photon-separation to enhance the spatial resolution of pulsed STED microscopy. Nanoscale. 11(4): 1754–1761. https://doi.org/10.1039/c8nr07485b
Liu, T., Kompa, J., Ling, J., Lardon, N., Zhang, Y., Chen, J., Reymond, L., Chen, P., Tran, M., Yang, Z., Zhang, H., Liu, Y., Pitsch, S., Zou, P., Wang, L., Johnsson, K. and Chen, Z. (2024) Gentle Rhodamines for Live-Cell Fluorescence Microscopy. ACS Cent Sci. 10(10): 1933-1944. https://doi.org/10.1021/acscentsci.4c00616
Article Information
Publication history
Received: Oct 4, 2024
Accepted: Dec 23, 2024
Available online: Jan 9, 2025
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© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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5,203 | https://bio-protocol.org/en/bpdetail?id=5203&type=0 | # Bio-Protocol Content
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This is an In Press version of the protocol that has not yet been assigned to an issue.
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Development of a Rapid and Efficient Protocol for Seed Germination and Seedling Establishment of Oryza coarctata
LB Lamis Berqdar
Mohamed A. Salem
JW Jian You Wang
AA Amer Alrudayan
MJ Muhammad Jamil
SA Salim Al-Babili
In Press, Available online: Jan 09, 2025
DOI: 10.21769/BioProtoc.5203 Views: 116
Reviewed by: Samik BhattacharyaArsheed Hussain Sheikh Anonymous reviewer(s)
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Abstract
Seed germination is a critical and challenging process in the propagation of Oryza coarctata, a wild halophytic rice species. This protocol outlines the seed germination procedure for O. coarctata. All steps required for optimal germination and seedling establishment of O. coarctata in both sterile and soil-based systems are described in detail. Additionally, the protocol includes an analysis of the primary hormones, abscisic acid (ABA) and gibberellin (GA), involved in regulating seed dormancy and germination.
Key features
• This protocol provides detailed instructions for germinating Oryza coarctata and improving its germination rate.
• The protocol enables the successful vegetative and reproductive growth of O. coarctata in soil.
• Additionally, the protocol outlines the analysis of hormones (abscisic acid and gibberellin) that play key roles in the germination process.
Keywords: Oryza coarctata Wild rice Germination Abscisic acid Gibberellin Seedlings
Graphical overview
Protocol for germination, seedling development, and hormone analysis of Oryza coarctata seeds. Step-by-step instructions include germinating seeds on filter papers or sterile growing medium; the established seedlings can be grown in controlled environments or a greenhouse. Extraction and analysis of endogenous key hormones offer insights into their roles in controlling dormancy and germination.
Background
Oryza coarctata is a perennial wild species of rice and the only halophytic species in the genus Oryza (family Poaceae) [1]. It has unique morphological features such as thicker waxy succulent leaves and salt glands, allowing the plant to survive under harsh environmental conditions such as high salinity and submersion [2]. Additionally, O. coarctata leaves have a unique leaf anatomical feature exhibited by C4 plants, the Kranz anatomy (enlarged bundle sheath cells encircling vascular bundles) [3,4]. O. coarctata is primarily found in coastal areas across Southeast Asian countries [1]. O. coarctata offers a wealth of genetic resources for rice breeding research, making it a good model system for investigating mechanisms of salinity and submersion tolerance in rice [5,6]. The germination rate of wild rice seeds is very low, posing significant challenges for research on its conservation. The low germination rate of wild rice seeds leads to inconsistent seedling emergence, causing a reduction in their yield [7]. Dormancy is a limiting factor for the breeding of wild rice, which is regulated by several plant hormones, mainly abscisic acid and gibberellin.
Biotic and abiotic stresses significantly challenge environmental stability and influence the survival of living organisms, including plants. For instance, salinity stress poses significant challenges to global agriculture, affecting production yield, which is essential for human consumption. In addition, salt stress affects around 6% of the world’s cultivable land, making it a significant threat to food security [8,9]. Salt accumulation has various effects, including reduced plant expansion, accelerated wilting, and disrupted photosynthesis. While changing climate intensifies these obstacles, addressing approaches to agriculture, such as genetic engineering to improve crop resilience in response to climatic stresses, is imperative [10,11]. Amongst various climatic stresses, salt concentration remarkably affects rice (Oryza sativa), one of the essential crops. Under these circumstances, the wild rice relative, O. coarctata, has become increasingly significant due to its resistance under salt stress compared to O. sativa. Moreover, cultivation efforts are directed toward producing saline-tolerant crops [12,13]. Therefore, the salt-tolerant characteristics of O. coarctata establish resources for cultivation programs intended to improve the adaptability of rice varieties to high salt levels, thus promoting food stability in regions affected by elevated salinity [4]. Additionally, O. coarctata is a promising candidate for neo-domestication.
Despite its potential as a promising crop for nutritional security, the germination of O. coarctata under experimental and laboratory conditions is challenging. Seed germination is controlled by phytohormones such as gibberellins (GAs) and abscisic acid (ABA) [14]. These hormones are also of particular significance given their role in the regulation of growth and development as well as responses to environmental stresses [15]. For instance, ABA controls plant development from germination to seed dormancy and modulates plant responses to challenging abiotic stresses [16,17]. Apart from their role in inducing seed germination, GAs modulate stem elongation, flower development, fruit ripening, and leaf senescence [18]. The quantitative analysis of these phytohormones is frequently required in both basic plant research and agricultural studies.
Taken together, developing a fast, reproducible, and efficient germination protocol for O. coarctata is crucial for both fundamental research and breeding programs. Establishing this protocol is critical for studying and investigating O. coarctata characteristics, such as biological traits and genetic features, to enhance crop resilience against elevated salt levels in multiple regions worldwide. In this protocol, we outline detailed steps for seed germination and seedling development of O. coarctata. Additionally, we determine the content of key hormones, offering new insights into their roles in controlling dormancy, germination, seedling establishment, and responses to environmental stresses.
Materials and reagents
Biological materials
1. Oryza coarctata seeds; original seeds were obtained from the International Rice Research Institute (IRRI), Philippines. For germination tests, seeds were harvested in our laboratory from plants that were grown at King Abdullah University of Science and Technology (KAUST) greenhouse facility
Reagents
1. Commercial bleach, sodium hypochlorite solution (NaClO) Sigma-Aldrich, CAS number: 7681-52-9)
2. Murashige and Skoog basal medium (MS) (Sigma-Aldrich, catalog number: M5519)
3. Sucrose (Sigma-Aldrich, CAS number: 57-50-1)
4. Agar (Sigma-Aldrich, CAS number: 9002-18-0)
5. LC-grade methanol (VWR, CAS number: 67-56-1)
6. LC-grade water (VWR, CAS number: 7732-18-5)
7. LC-grade acetonitrile (Fisher ChemicalTM, CAS number: 75-05-8)
8. Formic acid (Thermo ScientificTM, CAS number: 64-18-6)
9. Gibberellin A1 (GA1) (Sigma-Aldrich, CAS number: 545-97-1)
10. Abscisic acid (ABA) (Sigma-Aldrich, CAS number: 21293-29-8)
Solutions
1. 1% NaClO (100 mL) (see Recipes)
2. 0.5× MS media (100 mL) (see Recipes)
3. 50% acetonitrile:water (see Recipes)
4. Abscisic acid (ABA) stock solution (1 mg/mL) (see Recipes)
5. Gibberellin A1 (GA1) stock solution (1 mg/mL) (see Recipes)
6. Water containing 0.1% formic acid (1,000 mL) (see Recipes)
7. Acetonitrile containing 0.1% formic acid (1,000 mL) (see Recipes)
Recipes
1. 1% NaClO (100 mL)
Reagent Final concentration Quantity or Volume
NaClO (5% w/v) 1% (w/v) 20 mL
Water (ddH2O) - 100 mL*
*Note: Water should be added in increments to reach 100 mL final volume. This solution can be prepared and stored protected from light at room temperature for up to one month.
2. 0.5× MS media (100 mL)
Reagent Final concentration Quantity or Volume
Sucrose 2% (w/v) 2 g
MS powder 0.22% (w/v) 0.22 g
Water (ddH2O) n/a 100 mL*
Adjust pH to 5.8 with 3N HCl or 3N KOH. Add agar to 0.8% and autoclave at 121 °C for 15 min. The medium should be freshly prepared.
*Note: Water should be added in increments to reach 100 mL final volume.
3. 50% acetonitrile:water
Reagent Final concentration Quantity or Volume
LC-grade water 50% 50 mL
LC-grade acetonitrile 50% 50 mL
Total n/a 100 mL*
*Note: This solution can be prepared and stored protected from light at room temperature for up to three months.
4. Abscisic acid (ABA) stock solution (1 mg/mL)
Reagent Final concentration Quantity or Volume
Abscisic acid 1 mg/mL 1 mg
LC-grade methanol - 1 mL*
*Note: Working solution (10 µg/mL) from this stock can be prepared and stored protected from light at −20 °C for up to six months.
5. Gibberellin A1 (GA1) stock solution (1 mg/mL)
Reagent Final concentration Quantity or Volume
Gibberellin A1 1 mg/mL 1 mg
LC-grade methanol - 1 mL*
*Note: Working solution (10 µg/mL) from this stock can be prepared and stored protected from light at −20 °C for up to six months.
6. Water containing 0.1% formic acid (1,000 mL)
Reagent Final concentration Quantity or Volume
Formic acid 0.1% 1 mL
LC-grade water 100% Up to 1000 mL*
*Note: Water should be added in increments to reach 1000 mL final volume. This solution should be freshly prepared before analysis.
7. Acetonitrile containing 0.1% formic acid (1,000 mL)
Reagent Final concentration Quantity or Volume
Formic acid 0.1% 1 mL
LC-grade acetonitrile 100% Up to 1,000 mL*
*Note: Acetonitrile should be added in increments to reach 1,000 mL final solution.
This solution should be freshly prepared before analysis.
Laboratory supplies
1. Sterile deionized water, room temperature
2. 50 mL tubes (FalconTM, catalog number: 14-432-22)
3. Filter paper (Whatman®, Grade 1, model number: WHA1001090)
4. Parafilm (Fisher Scientific, catalog number: S37440)
5. Laminar flow hood (AIREGARD ES Horizontal Laminar Flow, model number: NU-201/E)
6. Petri dishes (FisherbrandTM, catalog number: AS4052)
7. Tissue culture boxes (e.g., Magenta Vessel with cover) (Sigma-Aldrich, model number: C0542)
8. 5″ diameter round black pots (with holes at the base for drainage)
9. Mixture of soil (Stender) and local sand (3:1)
10. 0.2 µm filter (Target2TM PTFE Syringe Filters, Thermo Scientific™, catalog number: F2504-4)
11. Steel beads (3.2 mm chrome-steel beads, BioSpec Products, Inc., catalog number: 11079132c)
12. 2 mL safe-lock microcentrifuge tubes (Eppendorf, catalog number: 0030123620)
13. 1.5 mL safe-lock microcentrifuge tubes (Eppendorf, catalog number: 0030123611)
14. Autosampler vials (LC screw-thread vials with caps and inserts, VWR®, catalog number: 82030-974A)
15. HPLC column (Hypersil GOLD C18 Selectivity, 150 × 4.6 mm; 3 µm; Thermo ScientificTM, catalog number: 25003-154630)
Equipment
1. pH meter (FisherbrandTM accumetTM AB150 pH benchtop meter, catalog number: 13-636-AB150)
2. 30 °C incubator (FisherbrandTM IsotempTM Incubator, catalog number: 15-103-0515)
3. Culture room/growth chamber (e.g., Conviron, BioChambers, or Percival) [with controlled LED light and humidity controller with humidity range from 60% to 90% relative humidity (RH) and temperature range from 10 °C to 40 °C]
4. Weighing balance (Sartorius M-Power Analytical Balance, model number: AZ214)
5. Tissue grinder (Mixer Mill, RETSCH®, model number: 20.746.0001)
6. Vortexer (VWR® Fixed Speed Vortex Mixer, catalog number: 10153-834)
7. Sonicator (Bransonic® Ultrasonic Cleaner, model: 5510)
8. Centrifuge (EppendorfTM 5424R Microcentrifuge, catalog number: 05-401-205)
9. Vacuum centrifugal concentrator (Eppendorf® Concentrator Plus, catalog number: 5305000568)
10. Liquid chromatography system (Thermo Scientific, model: VanquishTM Duo UHPLC System)
11. Mass spectrometer system (Thermo Scientific, model: TSQ AltisTM triple quadrupole)
Software and datasets
1. Statistical software for data analysis software (Microsoft Excel, Microsoft Corporation, v. 2410, 2024)
2. LC–MS/MS software (Xcalibur 4.1, Thermo Fisher Scientific Inc., USA)
Procedure
A. Seed germination
1. Before germination, remove the husks from the seeds.
2. Put the de-husked seeds in a 50 mL Falcon tube.
3. Proceed to sterilize them with 1% bleach solution on a vortex shaker (100 rpm) for 5 min.
4. Thoroughly rinse the seeds with 40 mL of sterile water at least five times. See General Note 1.
5. Prepare magenta boxes containing 0.5× MS media for section B or a Petri dish by placing two filter papers inside with 5 mL of sterile water for section C. For submerged germination, see General Note 2. The germination can be scored daily for one week. For dormant or non-germinated seeds, continue scoring for 12 days. The germination process requires a constant humid environment with temperature control for 7–10 days.
B. Growing O. coarctata in controlled environments
1. Place 5–8 disinfected seeds on magenta boxes (tissue culture boxes) containing 0.5× MS media.
2. Keep the boxes at 30 °C in the dark for 24–48 h.
3. Transfer the box containing germinated seeds to a controlled environment for growth.
4. Maintain a day/night temperature of 28/22 °C with a 12/12 h photoperiod, 200 µmol photons m−2 s−1, and 85% relative humidity (RH). The germination and seedling establishment can be scored for 7–10 days.
C. Growing O. coarctata in the greenhouse
1. Transfer 5–8 disinfected seeds onto the filter papers.
2. Seal the petri dish with parafilm to maintain seed moisture.
3. Incubate the Petri dish at 30 °C for 24–48 h in the dark.
4. Transfer the plates containing germinated seeds to a controlled environment for growth.
5. Maintain a day/night temperature of 28/22 °C with a 12/12 h photoperiod and 200 µmol photons m−2 s−1.
6. Fill 5″ diameter pots with a mixture of sand and soil (3:1).
7. Plant two seedlings per pot.
8. Transfer the pots containing seedlings to the greenhouse chamber. The temperature in our greenhouse ranges from 25 to 29 °C during the day and 22 to 25 °C at night. RH varies from 60% to 90%. The greenhouse is supplemented with artificial lights from 6 am to 7 pm. The entire reproductive growth is reached within three months.
9. After two weeks in the greenhouse, plants can also be retransferred to a shade house.
10. The plants can be irrigated on a daily basis and fertilized weekly with nitrogen, phosphorus, and potassium (NPK) fertilizer (20:20:20, Folicat Plantifol) at 2 g/L water.
D. Seed harvesting and storage
1. Seed collection: Harvest the seeds when they begin to turn brown, typically about two weeks after flowering.
2. Preparation: Place the collected seeds into tubes or suitable containers.
3. Storage: Store the seeds in a tightly sealed container at a cold temperature (around 4 °C) to preserve their viability. After the seed reaches maturity, wild rice goes into a dormant state. To break dormancy, seeds need to be kept in low temperatures for a few months.
E. Extraction and analysis of endogenous GA and ABA by LC/MS
1. Aliquot (50 mg) seeds in safe-lock 2.0 mL Eppendorf tubes.
2. Grind and homogenize seeds for 1 min at a frequency of 25 Hz.
3. Add 1.5 mL of LC-grade methanol to each tube.
4. Vortex samples at 1,000 rpm for 1 min.
5. Sonicate the samples at 40 kHz for 15 min.
6. Centrifuge samples at 10,000× g for 10 min at room temperature.
7. Transfer the supernatant to a new 1.5 mL Eppendorf tube.
8. Dry the supernatant by vacuum concentration for 1.5 h.
9. Resuspend the dry extracts in 120 µL of 50% acetonitrile:water followed by 1 min sonication at 40 kHz.
10. Filter the solution through a 0.22 µm filter into a LC glass vial.
11. Analyze samples by LC–MS/MS. See General Note 3.
12. Include standards for LC–MS/MS analysis in the running sequence. Dilute the standard hormones with methanol to 10 μg/mL (working solution) from the stock solution of 1 mg/mL ABA and the stock solution of 1 mg/mL GA1.
Data analysis
Germination rate estimation is based on scoring the percentage of germinated seeds every 24 h for 7 days. Use the following formula: Germination % = seeds germinated/total seeds × 100. Microsoft Excel or any other software can be used. Consider that radicle protrusion is the germination event.
The obtained LC/MS raw chromatograms from phytohormone analysis can be manually inspected using Xcalibur 4.1 (Thermo Fisher Scientific Inc., USA) or any other software. The retention time and peak areas can be recorded. For absolute quantification of endogenous hormones, use isotope-labeled standards as internal references during analysis [19].
Validation of protocol
In this protocol, we detail a rapid and efficient method for the germination of Oryza coarctata seeds. The protocol is validated through the figures and results demonstrating the functionality of the experimental setup. Using this method, radicle emergence was observed within 24–48 h and allowed us to obtain established seedlings in 10 days (Figure 1). Additionally, we analyzed the tolerance of O. coarctata seeds to submersion in water. We observed that seedling vigor was not affected.
Figure 1. Representative photographs of Oryza coarctata seeds at different days of germination initiation. Photographs were taken for dry seeds (A), 2 days after germination (B), and 10 days after sowing (C and D). Germination after submersion in water for 7 days is shown in (E).
Scoring the seed germination rate after the radical emergence, which is defined as the first sign of seed germination, revealed that around 40% of seeds successfully germinate after two days (Figure 2). Seeds are scored every day until seed germination rate reaches over 80% after five days.
Figure 2. Germination percentage of Oryza coarctata scored five days after sowing. Data are shown as mean ± SE (n = 10). In each replicate, 10 seeds were subjected to germination.
Optimizing the yield of O. coarctata depends on precise harvest timing, ideally during the green, milky stage (the stage when the grains are being filled) or approximately two weeks after flowering. The flowering period for O. coarctata typically spans under our growth conditions from December to January (Figure 3).
Figure 3. Representative photographs of Oryza coarctata plants vegetatively grown in the greenhouse. Photographs of two-week-old (A), one-month-old (B), and three-month-old (C–E) plants.
Simultaneous determination of GA and ABA from dry seeds was achieved by liquid chromatography–tandem mass spectrometry (LC–MS/MS) method. The extraction method with a single solvent (methanol) allows the detection of both hormones without the solid–phase extraction (SPE) pretreatment method. The chromatographic separation was carried out on a reversed-phase (C18) column, using acetonitrile/water containing 0.1% formic acid as mobile phases. Both phytohormones were eluted within 10 min. The full mass spectrum of ABA showed that [M−H]− ion was the most intensive at m/z 263.20 with a retention time of 9.67 min (Figure 4A). Multiple reaction monitoring (MRM) transitions (precursor ion/ product ion) are 263.20/153.10 and 263.20/219.10 (Figure 4B).
Figure 4. Typical LC chromatogram (A) and negative ion MS/MS spectrum (B) from LC–ESI-MS/MS analysis of abscisic acid (ABA) standard solution (upper panel) and Oryza coarctata seed extract (lower panel). The MRM transition for ABA, 263.20→ 219.10, 153.10. The chemical structure of ABA is shown.
The full mass spectrum of GA1 showed that the [M−H]− ion was the most intensive at m/z 347.20 with a retention time of 8.47 min (Figure 5A). MRM transitions are 347.20/259.10 and 347.20/273.00 (Figure 5B).
In conclusion, we have developed an efficient protocol for seed germination and seedling development of the wild rice species (Oryza coarctata) under sterile conditions for tissue culture and other agricultural applications.
Figure 5. Typical LC chromatogram (A) and negative ion MS/MS spectrum (B) from LC–ESI-MS/MS analysis of gibberellin A1 (GA1) standard solution (upper panel) and Oryza coarctata seed extract (lower panel). The MRM transitions for GA1, 347.20→ 273.00, 259.10. The chemical structure of GA1 is shown.
General notes and troubleshooting
General notes
1. Perform steps in a laminar flow hood under sterile conditions.
2. For the submerged germination, place the seeds in a sterile tube filled to 90% capacity with sterile water, positioning them diagonally. Incubate the tubes in a 30 °C incubator for 24–48 h in the dark. Transfer the tubes containing germinated seeds to a controlled environment for growth. Maintain a day/night temperature of 28/22 °C with a 12/12 h photoperiod and 200 µmol photons m−2 s−1.
3. The LC–MS analysis was performed following our previous studies [20,21]. Chromatographic separation was carried out on a C18 HPLC column maintained at 35 °C. Mobile phases consist of water containing 0.1% formic acid (A) and acetonitrile containing 0.1% formic acid (B). The gradient program was as follows: 0–10 min, 15% B to 100% B; 10–15 min, 100% B; 15–17 min, 15% B at 0.5 mL/min flow rate. The MS parameters were as follows: negative ion, 3000 V; sheath gas, 40 Arb; aux gas, 15 Arb; collision energy of 20 eV; ion transfer tube temperature, 350 °C; vaporizer temperature, 350 °C; cycle time, 1 s; Q1/Q3 resolution (FWHM), 0.4; CID gas (mTorr), 2; and chromatographic peak width (s), 6.
Troubleshooting
Problem 1: No germinations after three days.
Possible cause: Seeds are dormant.
Solution: Wait for at least seven days. Sometimes, seeds need more time to start germination (up to 10 days in the dark).
Problem 2: Fungal growth in culture boxes.
Possible cause: Contamination during seed sterilization or media preparation.
Solution: Discard the culture boxes and begin again under controlled sterile conditions. Perform steps in a laminar flow hood under sterile conditions.
Problem 3: No detection of ABA or GA signal from tissues but clear signal for standard.
Possible cause: The endogenous level is below the detection level.
Solution: Start extraction with ~100 mg of dry tissues.
Acknowledgments
This work was supported by Neo-domestication funding (CDA-neodom) and baseline funding given to S. A-B from King Abdullah University of Science and Technology (KAUST).
Competing interests
The authors declare that they have no competing interests.
References
Chowrasia, S., Rawal, H. C., Mazumder, A., Gaikwad, K., Sharma, T. R., Singh, N. K. and Mondal, T. K. (2018). Oryza coarctata Roxb. Mondal, T. K. and Henry, R. J. (Eds.) The Wild Oryza Genomes. Springer International Publishing. 87–104. https://doi.org/10.1007/978-3-319-71997-9_8
Bansal, J., Gupta, K., Rajkumar, M. S., Garg, R. and Jain, M. (2020). Draft genome and transcriptome analyses of halophyte rice Oryza coarctata provide resources for salinity and submergence stress response factors. Physiol Plant. 173(4): 1309–1322. https://doi.org/10.1111/ppl.13284
Zhao, H., Wang, W., Yang, Y., Wang, Z., Sun, J., Yuan, K., Rabbi, S. M. H. A., Khanam, M., Kabir, M. S., Seraj, Z. I., et al. (2023). A high-quality chromosome-level wild rice genome of Oryza coarctata. Sci Data. 10(1): 701. https://doi.org/10.1038/s41597-023-02594-1
Chowrasia, S., Nishad, J., Pandey, R. and Mondal, T. K. (2021). Oryza coarctata is a triploid plant with initial events of C4 photosynthesis evolution. Plant Sci. 308: 110878. https://doi.org/10.1016/j.plantsci.2021.110878
Mondal, T. K., Rawal, H. C., Gaikwad, K., Sharma, T. R. and Singh, N. K. (2017). First de novo draft genome sequence of Oryza coarctata, the only halophytic species in the genus Oryza. F1000Research. 6: 1750. https://doi.org/10.12688/f1000research.12414.1
Bansal, J., Gupta, K., Rajkumar, M. S., Garg, R. and Jain, M. (2020). Draft genome and transcriptome analyses of halophyte rice Oryza coarctata provide resources for salinity and submergence stress response factors. Physiol Plant. 173(4): 1309–1322. https://doi.org/10.1111/ppl.13284
Yao, Q., Zheng, X., Zhou, G. and Zhang, J. (2024). SGR-YOLO: a method for detecting seed germination rate in wild rice. Front Plant Sci. 14: e1305081. https://doi.org/10.3389/fpls.2023.1305081
Rodríguez Coca, L. I., García González, M. T., Gil Unday, Z., Jiménez Hernández, J., Rodríguez Jáuregui, M. M. and Fernández Cancio, Y. (2023). Effects of Sodium Salinity on Rice (Oryza sativa L.) Cultivation: A Review. Sustainability. 15(3): 1804. https://doi.org/10.3390/su15031804
Teshome, D. T., Zharare, G. E. and Naidoo, S. (2020). The Threat of the Combined Effect of Biotic and Abiotic Stress Factors in Forestry Under a Changing Climate. Front Plant Sci. 11: e601009. https://doi.org/10.3389/fpls.2020.601009
Liu, X., Feike, T., Shao, L., Sun, H., Chen, S. and Zhang, X. (2016). Effects of saline irrigation on soil salt accumulation and grain yield in the winter wheat-summer maize double cropping system in the low plain of North China. J Integr Agric. 15(12): 2886–2898. https://doi.org/10.1016/s2095-3119(15)61328-4
Khan, M. S., Akther, T., Mubarak Ali, D. and Hemalatha, S. (2019). An investigation on the role of salicylic acid alleviate the saline stress in rice crop (Oryza sativa (L)). Biocatal Agric Biotechnol. 18: 101027. https://doi.org/10.1016/j.bcab.2019.101027
Tamanna, N., Mojumder, A., Azim, T., Iqbal, M. I., Alam, M. N. U., Rahman, A. and Seraj, Z. I. (2024). Comparative metabolite profiling of salt sensitive Oryza sativa and the halophytic wild rice Oryza coarctata under salt stress. Plant-Environ Interact. 5(3): e10155. https://doi.org/10.1002/pei3.10155
Mukherjee, S., Mukherjee, A., Das, P., Bandyopadhyay, S., Chattopadhyay, D., Chatterjee, J. and Majumder, A. L. (2021). A salt‐tolerant chloroplastic FBPase from Oryza coarctata confers improved photosynthesis with higher yield and multi‐stress tolerance to indica rice. Plant Cell, Tissue Organ Cult. 145(3): 561–578. https://doi.org/10.1007/s11240-021-02026-1
Zhang, Y., Berman, A. and Shani, E. (2023). Plant Hormone Transport and Localization: Signaling Molecules on the Move. Annu Rev Plant Biol. 74(1): 453–479. https://doi.org/10.1146/annurev-arplant-070722-015329
Waadt, R., Seller, C. A., Hsu, P. K., Takahashi, Y., Munemasa, S. and Schroeder, J. I. (2022). Plant hormone regulation of abiotic stress responses. Nat Rev Mol Cell Biol. 23(10): 680–694. https://doi.org/10.1038/s41580-022-00479-6
Wang, J. Y., Lin, P. Y. and Al-Babili, S. (2021). On the biosynthesis and evolution of apocarotenoid plant growth regulators. Semin Cell Dev Biol. 109: 3–11. https://doi.org/10.1016/j.semcdb.2020.07.007
Jamil, M., Alagoz, Y., Wang, J. Y., Chen, G. E., Berqdar, L., Kharbatia, N. M., Moreno, J. C., Kuijer, H. N. J. and Al‐Babili, S. (2024). Abscisic acid inhibits germination of Striga seeds and is released by them likely as a rhizospheric signal supporting host infestation. Plant J. 117(5): 1305–1316. https://doi.org/10.1111/tpj.16610
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Salem, M. A., Yoshida, T., Perez de Souza, L., Alseekh, S., Bajdzienko, K., Fernie, A. R. and Giavalisco, P. (2020). An improved extraction method enables the comprehensive analysis of lipids, proteins, metabolites and phytohormones from a single sample of leaf tissue under water‐deficit stress. Plant J. 103(4): 1614–1632. https://doi.org/10.1111/tpj.14800
Wang, J. Y., Alseekh, S., Xiao, T., Ablazov, A., Perez de Souza, L., Fiorilli, V., Anggarani, M., Lin, P. Y., Votta, C., Novero, M., et al. (2021). Multi-omics approaches explain the growth-promoting effect of the apocarotenoid growth regulator zaxinone in rice. Commun Biol. 4(1): 1222. https://doi.org/10.1038/s42003-021-02740-8
Chen, G. T., Wang, J. Y., Votta, C., Braguy, J., Jamil, M., Kirschner, G. K., Fiorilli, V., Berqdar, L., Balakrishna, A., Blilou, I., et al. (2023). Disruption of the rice 4-DEOXYOROBANCHOL HYDROXYLASE unravels specific functions of canonical strigolactones. Proc Natl Acad Sci USA. 120(42): e2306263120. https://doi.org/10.1073/pnas.2306263120
Article Information
Publication history
Received: Oct 15, 2024
Accepted: Dec 23, 2024
Available online: Jan 9, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Peer-reviewed
Vegetative Propagation of Cannabis sativa and Resin Obtained From its Female Inflorescences
SD Sebastián D´Ippolito
ML Marina Landaburu
MB María E. Vozza Berardo
MV María D. Villamonte
JM Julieta R. Mendieta
DN Débora Nercessian
SC Silvana L. Colman
In Press, Available online: Jan 19, 2025
DOI: 10.21769/BioProtoc.5204 Views: 27
Reviewed by: Noelia ForesiSimab Kanwal Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Journal of Ethnopharmacology Jan 2024
Abstract
Cannabis (Cannabis sativa L.) derivatives are of great importance in the medical, cosmetic, and pharmaceutical industries. This relevance is mainly due to the active principles (cannabinoids) found mainly in the trichomes of the female inflorescences. One of the most commonly used methods to propagate cannabis is by vegetative stem cuttings. This low-cost technique produces genetically uniform plants, ensuring consistent growth rates and cannabinoid production. The extraction of cannabinoids and other active compounds from the resin of the flowers is the main limitation of cannabis processing. Here, we describe a step-by-step protocol for propagating female cannabis plants from vegetative stem cuttings, inducing flower development, and obtaining high-quality cannabinoid-enriched resin.
Key features
• The propagation of cannabis is done by vegetative cuttings.
• It takes 4–5 months to cultivate the plant and obtain female inflorescences.
• Resin is the end product of the alcoholic extraction of flowers, and the obtained resin is a full-spectrum mixture of active compounds.
Keywords: Cannabis sativa Vegetative propagation Stem cutting Inflorescences Resin Cannabinoids
Graphical overview
Main steps for the vegetative propagation of cannabis and extraction of resin from female inflorescences. The figure highlights the characteristics of the cutting and its arrangement in the corresponding tray. Additionally, it shows female inflorescences both in vivo and after harvesting and subsequent drying. From this point, different resin yields can be obtained depending on the protocol used.
Background
Cannabis sativa L. is an annual dicotyledonous herbaceous plant of the Cannabaceae family [1]. Nowadays, the female plant is mainly used in the medical, cosmetic, and pharmaceutical industries. The development of new products in these health-related fields is mainly due to the therapeutic effects associated with various cannabinoids [2]. These compounds are present in trichomes, glandular structures located mainly in female inflorescences [3,4]. Currently, several subclasses of cannabinoids have been identified and categorized into 11 families, each containing a variable number of compounds. The best-known cannabinoids include tetrahydrocannabinol, cannabidiol, cannabinol, cannabichromene, cannabicyclol, and cannabigerol. In addition, there is a wide variety of terpenes, sugars, steroids, flavonoids, and nitrogenous compounds that give flavor and aroma to different strains and have synergistic effects with cannabinoids [5]. The ratios of active compounds in flowers are influenced by a number of factors, including plant genetics, growing and storage conditions, maturity at harvest, and the methods used to process and formulate the material [6].
To maintain the genetic stability of a plant with a specific chemical profile, asexual propagation by vegetative stem cutting is a very suitable method [7]. In this technique, a section of the plant is taken to create a clone that is genetically identical to the parent plant. It is then placed on a moist substrate to allow the roots to develop. When the root system is thriving, it is transplanted to its final location where the plant will grow vegetatively until it reaches flowering and subsequent cannabinoid production. Although this technique is simple, the success of this method depends on many variables such as plant genetics, explant characteristics, hormones, and growing conditions such as light, humidity, and temperature [8].
Once harvested, flowers should be trimmed and then dried (in the absence of sunlight to prevent photochemical transformation) before beginning extraction of the active compounds. Cannabinoids are located in the trichomes on the surface of the bud, where the concentration gradient between the solvent and the trichome surface acts as the primary barrier to chemical transfer. Ethanol is the solvent of choice because of its solubility, relatively low price, and more importantly, its boiling point (for recovery purposes). For medical or dietary applications, ethanol is preferred due to its lower toxicity [9]. After extraction, the solvent is removed to yield a resin consisting of cannabinoids, terpenes, and other accompanying compounds. Depending on the final use of the resin, additional steps for purification and/or isolation are required. In this paper, we specifically detail the procedure to effectively generate cuttings and then list the steps to achieve an efficient resin yield from an alcoholic extraction of cannabinoids.
Materials and reagents
Biological materials
1. Cannabis sativa L. plants
Reagents
1. Substrate GrowMix Multipro (Terrafertil)
2. Substrate enriched with worm castings, peat, river sand, pine needles, and perlite (Pura Pacha)
3. Root promoter, alpha naphthalene acetic acid 0.1 g/100 mL (Fertifox)
4. Ethanol 96% and 70% (Porta, CAS 64-17-5)
5. Sodium hypochlorite 30% (Odex, CAS 7681-52-9)
6. Fertilizer NPK 5-9-12 Top Bloom (Top Crop)
Laboratory supplies
1. Bootstrap farmer 72-cell tray (Carluccio)
2. Transparent plastic box 60 cm × 50 cm × 30 cm (Colombraro, catalog number: 257)
3. Pots (1 and 5 L) (Carluccio)
4. Falcon tubes 50 mL (Deltalab, catalog number: 429931)
5. Eppendorf tubes 1.5 mL (Deltalab, catalog number: 4092.2AM)
6. Pipette tips 1,000 μL (Deltalab, catalog number: 200070 309)
7. Airtight bags (Ziploc)
8. Beaker 250 mL (Borosil, catalog number: 1000D21)
9. Laboratory gloves (Fisher Scientific, catalog number: 19-130-1597B)
10. Qualitative filter paper Grade 1 (Whatman, catalog number: WHA1001125)
11. Scissors
Equipment
1. Cultivation room at 25 °C, 18/6 h light/dark photoperiod for vegetative stages and 12/12 h for floral stages. Led hell panels 400 W (Led Osram HyperRED 660 nm, Led duris E2836 4000K and 3000K, Cri80, Drivers Philips)
2. Homogenizer (IKA, model: T10 basic ULTRA-TURRAX)
3. PIPETMAN P1000L 100–1,000 μL (Gilson, model: F167370)
4. Analytical balance (Shimadzu, model: AUY220)
5. Automatic environmental SpeedVac system (Savant, model: AES1010)
6. Rotary evaporator (Dragon Lab, model: RE-100 PRO)
7. Centrifuge (Thermo Scientific, model: Sorvall ST16R)
Procedure
A. Obtaining cuttings
To obtain cuttings, start with a healthy Cannabis sativa L. plant in the vegetative phase of growth, with green branches. The plant must be free of visible pathogens and without symptoms of nutrient deficiency or excess (Figure 1).
Figure 1. Vegetative propagation of cannabis by stem cuttings procedure. The diagram illustrates the process of taking a cutting from a mother plant in the vegetative stage, highlighting the area where the rooting hormone is applied. The cuttings are then placed in a tray with substrate within a wet chamber, where they remain until visible roots form.
1. Select a green branch with at least three nodes and 6 cm in length.
2. Sterilize a pair of scissors with 70% alcohol.
3. Make a 45° cut just below the third node.
4. Completely remove the petiole and the leaf that emerges from the third node by making a cut through the petiole.
5. Remove a portion of the leaf tip (50%) of fully expanded leaves to reduce water loss through evaporation.
6. Submerge the stem of the cutting in tap water while making new cuttings.
7. Apply a rooting agent to the basal area of the cutting containing the third node.
B. Substrate preparation
Once a good-quality substrate is available, follow these steps:
1. Clean a tray with the required number of wells with 30% sodium hypochlorite and rinse with tap water.
2. Fill the required number of wells with substrate.
3. Moisten the substrate with potable water.
4. Make a hole in the center of the substrate.
5. Insert the cut into the substrate, making sure the third knot is below the substrate surface.
6. Gently press down with your fingertips to secure the cut.
7. Add more substrate until it reaches the edge of the tray.
8. Place the tray in a transparent plastic box or tub.
9. Close the lid of the transparent plastic box to create a wet chamber.
10. Place the “wet chamber” with cuttings in the cultivation room to allow root development.
C. Root development and growth of cutting
The rooting conditions should be as follows: 18/6 h (light/dark) photoperiod, light intensity 50 W/m2, temperature 24 °C, relative humidity 80%–100% inside the humid chamber. After three days, press the substrate with your fingertip to check if it remains moist. If the substrate is dry, add potable water until moisture is restored. After 14 days, check for the presence of roots. These can be visualized by observing the drainage surface of the seed tray. Alternatively, remove the block of substrate from the germination tray and observe root development in the substrate directly. Once the roots have developed, transplant the cutting into a 1 L pot pre-filled with organic substrate (Pura Pacha) to facilitate root expansion and growth. This process should be repeated with larger pots as needed. Rooted cuttings should be watered every two days and kept in vegetative growth under the following conditions: 18/6 h (light/dark) photoperiod, light intensity 100 W/m2, temperature 24 °C, relative humidity 75%, and continuous airflow. When the plants have reached an adequate vegetative development (3–5 weeks), transplant them to a 5 L pot; the flowering stage begins.
D. Flowering plants
To initiate the flowering stage, switch to a photoperiod of 12/12 h light/dark and maintain the following conditions: light intensity 200 W/m2, temperature 24 °C, relative humidity 60%–70%, and continuous airflow. Within a period of 7–14 days, the plants will begin to develop sexual characteristics or flowers. The development and maturation of the female flowers will take approximately two to three months, depending on the variety of cannabis.
Throughout this process, the plants should be watered with potable water and the substrate should be kept at a pH around 6 with adequate nutrients (NPK 5-9-12), free of pesticides or heavy metal residues. It is important to have airflow around the leaf surface by installing fans in the growing environment. Good air circulation also helps prevent the establishment of plant diseases.
E. Harvesting, drying, and storing flowers
At the end of the growing season, when the pistils begin to turn brown, label and harvest the plants. Cut the main stem at the bottom and the main branches if the plant has developed a large biomass. Remove larger leaves to allow airflow between the branches, then hang the plants/branches upside down to dry in the dark for 10 days. Then, separate the female flowers from the plants with scissors and manually remove the remaining leaves between the flowers. Wear gloves when handling the plant.
At this point, it is necessary to have an additional room or drying tent. This room should be kept dark and cool to prevent contamination of the flowers with fungi or other microorganisms. Once the flowers are dry, store them in airtight bags and place them in darkness. Open them for 1 min during the first 10 days to further remove any remaining moisture. The harvested material must be protected from air, temperature, and light to prevent processes such as oxidation and decarboxylation of cannabinoids. These dried flowers are now ready to be processed for resin extraction as described below.
F. Obtaining resin from female flowers
The most appropriate extraction method should be selected based on the characteristics of the desired product. Conventional extraction using organic solvents involves macerating the plant material and then removing the solvent under reduced pressure. It is important to exercise caution during extraction, as the constituents of the original plant compounds are exposed to heat, air, and pressure, which can result in chemical changes (Figure 2).
Figure 2. Procedure for resin extraction from female inflorescences. The procedure for working with small volumes (red arrows) involves the use of a centrifuge and SpeedVac equipment. For larger extraction volumes (blue arrows), the extract is decanted prior to filtration, and the solvent is removed using a rotary evaporator.
1. Weigh the dried female flowers on a precision balance.
2. Place the flowers in a sterilized container and add cold 96% ethanol at a 1/15 w/v ratio.
3. Process the samples by mechanical disruption using a homogenizer until a fine suspension is formed.
4. Separate the plant material from the liquid phase by centrifugation at 800× g for 5 min in Falcon tubes or allow it to decant at room temperature.
5. Retain the liquid phase, collect the solid material, and repeat steps F2–4.
6. Filter the liquid phase through grade 1 filter paper and save the filtered extract.
7. Remove solvent under vacuum. Depending on the volume obtained, different equipment may be required. For evaporating large volumes, a rotary evaporator may be required under the following conditions: 40 °C, 60× g, and 20 MPa for the vacuum pump until ethanol recovery is complete. For small volumes, the sample can be evaporated using a SpeedVac concentrator under the following conditions: radiant cover off, cryopumping, and low drying rate. Samples should be fractionated into pre-weighed Eppendorf tubes. For example, 1 mL of extract should be evaporated for approximately 4 h.
8. Store the obtained resin at -20 °C.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article:
• Vozza Berardo et al. [10]. Antifungal and antibacterial activities of Cannabis sativa L. resins. J Ethnopharmacology 318:116839. https://doi.org/10.1016/j.jep.2023.116839
General notes and troubleshooting
General notes
1. High relative humidity and adequate temperature (22–26 °C) are critical for successful explanting.
2. If no roots have appeared by day 15, discard the plants.
3. This resin can also be suspended in vegetable oil to obtain cannabis oil, a highly useful product in the medical, cosmetic, and pharmaceutical industries.
4. This protocol is also applicable to other research objectives (e.g., in vitro detached leaf/fruit inoculation) and flowering plants with appropriate modifications.
Troubleshooting
Problem 1: There is no root development.
Possible cause: The temperature in the chamber is too low. The stem is too thick.
Solution: Keep the temperature inside the chamber between 22 and 26 °C. Use green branches.
Problem 2: The cutting dries up.
Possible cause: The humidity inside the chamber is not adequate.
Solution: Keep the humidity inside the chamber between 80% and 100%.
Problem 3: The cutting rots before developing roots.
Possible cause: The substrate is too wet.
Solution: Be careful when wetting the substrate to avoid overdoing it.
Problem 4: There is remaining solvent in the resin.
Possible cause: The evaporation time was not enough.
Solution: Evaporate until no visible traces of solvent remain in the resin.
Acknowledgments
We are grateful to the following science and technology entities for technical support: Universidad Nacional de Mar del Plata and Comisión de Investigaciones Científicas.
This paper was supported by Agencia Nacional de Investigaciones Científicas y Técnicas (ANPCyT, PICT Serie A 03535) and Consejo Nacional de Investigaciones Científicas y Técnicas (CONICET) PIP 0422.
This protocol was described and validated in the original research article Vozza Berardo et al. [10]. Antifungal and antibacterial activities of Cannabis sativa L. resins. J Ethnopharmacology 318:116839.
Competing interests
The authors declare no competing interests.
References
Bremer, K. (1998). An Ordinal Classification for the Families of Flowering Plants. Ann Mo Bot Gard. 85(4): 531.
Abot, A., Bonnafous, C., Touchard, F., Thibault, F., Chocinski-Arnault, L., Lemoine, R. and Dédaldéchamp, F. (2013). Effects of cultural conditions on the hemp (Cannabis sativa) phloem fibres: Biological development and mechanical properties. J Compos Mater. 47(8): 1067–1077.
Schilling, S., Dowling, C. A., Shi, J., Ryan, L., Hunt, D., OReilly, E., Perry, A. S., Kinnane, O., McCabe, P. F., Melzer, R., et al. (2020). The Cream of the Crop: Biology, Breeding and Applications of Cannabis sativa. 10.22541/au.160139712.25104053/v2
Aizpurua-Olaizola, O., Soydaner, U., Öztürk, E., Schibano, D., Simsir, Y., Navarro, P., Etxebarria, N. and Usobiaga, A. (2016). Evolution of the Cannabinoid and Terpene Content during the Growth of Cannabis sativa Plants from Different Chemotypes. J Nat Prod. 79(2): 324–331.
Chandra, S., Lata, H., Mehmedic, Z., Khan, I. A. and ElSohly, M. A. (2015). Light dependence of photosynthesis and water vapor exchange characteristics in different high Δ9-THC yielding varieties of Cannabis sativa L. J Appl Res Med Aromat Plants. 2(2): 39–47.
Potter, D. J. (2014). A review of the cultivation and processing of cannabis (Cannabis sativa L.) for production of prescription medicines in the UK. Drug Test Anal. 6: 31–38.
de Meijer, E. P. M., Bagatta, M., Carboni, A., Crucitti, P., Moliterni, V. M. C., Ranalli, P. and Mandolino, G. (2003). The Inheritance of Chemical Phenotype in Cannabis sativa L. Genetics. 163(1): 335–346.
Caplan, D., Stemeroff, J., Dixon, M. and Zheng, Y. (2018). Vegetative propagation of cannabis by stem cuttings: effects of leaf number, cutting position, rooting hormone, and leaf tip removal. Can J Plant Sci. 98(5): 1126–1132.
Valizadehderakhshan, M., Shahbazi, A., Kazem-Rostami, M., Todd, M. S., Bhowmik, A. and Wang, L. (2021). Extraction of Cannabinoids from Cannabis sativa L. (Hemp)—Review. Agriculture. 11(5): 384.
Vozza Berardo, M. E., Mendieta, J. R., Villamonte, M. D., Colman, S. L. and Nercessian, D. (2024). Antifungal and antibacterial activities of Cannabis sativa L. resins. J Ethnopharmacol. 318: 116839.
Article Information
Publication history
Received: Sep 14, 2024
Accepted: Dec 16, 2024
Available online: Jan 19, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
How to cite
Category
Plant Science > Plant biochemistry > Metabolite
Biological Sciences > Biological techniques
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The authors used this protocol in Journal of Molecular Biology Mar 2024
Abstract
Cellular communication relies on the intricate interplay of signaling molecules, which come together to form the cell–cell interaction (CCI) network that orchestrates tissue behavior. Researchers have shown that shallow neural networks can effectively reconstruct the CCI from the abundant molecular data captured in spatial transcriptomics (ST). However, in scenarios characterized by sparse connections and excessive noise within the CCI, shallow networks are often susceptible to inaccuracies, leading to suboptimal reconstruction outcomes. To achieve a more comprehensive and precise CCI reconstruction, we propose a novel method called triple-enhancement-based graph neural network (TENET). The TENET framework has been implemented and evaluated on both real and synthetic ST datasets. This protocol primarily introduces our network architecture and its implementation.
Key features
• Cell–cell reconstruction network using ST data.
• To facilitate the implementation of a more holistic CCI, we incorporate diverse CCI modalities into consideration.
• To further enrich the input information, the downstream gene regulatory network (GRN) is also incorporated as an input to the network.
• The network architecture considers global and local cellular and genetic features rather than solely leveraging the graph neural network (GNN) to model such information.
Keywords: Cell–cell interaction network (CCI) reconstruction Gene regulatory network (GRN) Spatial transcriptomics (ST) data Graph neural network (GNN) Attention mechanism
Graphical overview
Graphical abstract of TENET, including (a) the knowledge graph preparation on both cell and gene levels and (b) the network architecture.
Background
Understanding cellular communication is crucial for constructing a cell–cell interaction network (CCI), which allows researchers to investigate the roles of different cells in biological processes and diseases. A common method for analyzing CCI involves studying the interactions between secreted ligands and their corresponding receptors (LR pairings), as these interactions are essential for signal transmission.
However, CCIs also occur through direct cell–cell contact, the extracellular matrix (ECM), and the secretion of signaling molecules [1]. Focusing solely on LR pairings overlooks the spatial context in which these interactions occur. With the advent of spatial transcriptomics (ST) data, incorporating spatial information can help address this limitation. Several methods for reconstructing CCI using ST data have been developed. For instance, Giotto [2] constructed a spatial grid based on cell coordinates to model proximal interactions but did not account for distal interactions. MISTy [3] utilizes multiple perspectives (intrinsic, local, tissue) to enhance cell knowledge for CCI inference. DeepLinc [4] employs a graph neural network (GNN) [5] to create a spatial proximity graph, using KNN [6] to identify both proximal and distal interactions. Despite these advances, modeling ST data solely at the cell level can lead to false positives and negatives due to a lack of downstream information. Incorporating downstream data, such as intracellular signaling, gene regulation, and protein changes can provide crucial insights for understanding actual CCI; CLARIFY [7] combines cellular and genetic information to build knowledge graphs for GNN inputs, effectively capturing structural features important for CCI inference. However, solely using GNNs is limiting, as they often aggregate local information, hindering the capture of global context. GNNs also struggle with similar nodes, which may lead to a loss of features’ specificity.
To overcome these challenges, we propose a novel approach called triple-enhancement-based graph neural network (TENET), designed as a comprehensive architecture for CCI reconstruction. TENET employs a graph convolution network (GCN) [8] backbone to extract global features from the data at multiple resolutions, generating robust latent feature embeddings. These embeddings then undergo a triple-enhancement mechanism that restores cell specificity that may have been lost during the GCN's global aggregation. The first enhancement mitigates the GCN's over-smoothing problem, while the secondary and tertiary enhancements refine the embeddings both locally and globally. Finally, a denoising loss function is introduced to tackle the challenges posed by noisy, low-quality data often encountered in small molecule experiments. This multi-scale enhancement strategy enables TENET to accurately and robustly reconstruct cell–cell interactions, addressing the limitations of previous methods and opening new avenues for researchers to explore the complexities of CCI.
Software and datasets
Software
1. 2.90 GHz Intel i7-10700F CPU and NVIDIA A100 graphics card; nvcc: NVIDIA (R) Cuda compiler driver; Copyright 2005–2024 NVIDIA Corporation; Built on Tue_Feb_27_16:19:38_PST_2024; Cuda compilation tools, release 11.7, V11.7.99; Build cuda_11.7.r11.7/compiler.33961263_0
Input data
1. Three ST datasets are utilized, namely seqFISH, MERFISH, and scMultiSim [9], which can be acquired at https://bitbucket.org/qzhu/smfish-hmrf/src/master/, https://datadryad.org/stash/dataset/10.5061/dryad.8t8s248, and https://github.com/ZhangLabGT/scMultiSim. The hyperlinks direct to the datasets utilized in TENET. The specific analysis of the datasets is mentioned in the following sections.
Procedure
The procedure includes two parts: the input preparation in section A and the network architecture in section B. The inputs of the network are the knowledge graphs from the cell and gene levels. Graphs encapsulate both the nodal feature representation V as well as the corresponding adjacency matrix (edge list) Adj. In section A, we elucidate the respective construction for the cell-level graph and the gene-level graph, denoted as Gc and Gg, respectively. In section B, we present the corresponding operations to the constructed knowledge graphs. Table 1 lists the frequently used symbols in sections A and B, improving overall readability.
Table 1. Frequently used symbols
Symbol Explanation
Gc(g) =(Vc(g) ,Adjc(g)) Input cell (gene) knowledge graph, containing nodes with features and the adjacency matrix.
Vc(g) ∈R(Nc(g) ×Fc(g)) Nc(g) number of nodes with Fc(g) feature dimensions.
Adjc(g) ∈R(Nc(g) ×Nc(g)) Adjacency matrix representing the establishment of interactions among nodes.
Kp The numbers of cells' indexes denoting the establishment of proximal interaction.
Adjp∈R(Nc×Nc) The proximal interaction adjacency matrix.
Kir The numbers of cells' indexes denoting the establishment of intermediate range interaction.
Adjir∈R(Nc×Nc) The intermediate range interaction adjacency matrix.
Zc(g) ∈R(Nc(g) × Fc(g)) Latent feature embeddings after the input graph Gc(g) traverse through the GCN encoder.
Z(c+g)∈R(Nc×F(c+g)) An integration of Zc and Zg.
Z'(c+g)∈R(Nc×F'(c+g)), Z'g∈R(Nc×F'g) Enhancing Z(c+g), Zg using CECB.
Z''(c+g)∈R(Nc×F''(c+g)), Z'g∈R(Nc×F''g) Enhancing Z(c+g)' and Zg' using EEB and SEB, respectively.
Z'''(c+g)∈R(Nc×F'''(c+g)), Z'g∈R(Nc×F'''g) Integration of the previous result {Z(c+g), Z(c+g,)' Z(c+g)'''},{Zg, Z(g,)' Zg'''}.
A. Input preparation
The construction of Gc and Gg is described in subsections a and b, respectively.
1. Cell-level graph construction
From the aforementioned links, there are four .csv files corresponding to each dataset. These files encompass the spatial coordinates of individual cells, the cell type classifications, the gene expression count matrices for each cell, and the adjacency matrix delineating cellular connectivity. Except for the adjacency matrix file, the other three files are encapsulated into a data frame, representing the cells' features. Figure 1 depicts the cell features organized into a data frame format; using the seqFISH dataset as an example, there are Nc number of cells with Fc number of features, denoted as Vc∈R(Nc×Fc). In this feature representation, the blue overlay delineates the spatial coordinates of individual cells, the green overlay denotes the cell type classifications, and the red overlay represents the gene expression matrix. (Due to space limitation, the red overlay is a partial visualization of the gene expression profile.) To build a cell-level adjacency matrix Adjc, our procedure is not simply to use the provided adjacency matrix but to simulate actual cell communication in the real biological world by integrating the three interaction modes into consideration. These three secretion modes exhibit a different range of interactions, where the cell–cell direct contact is defined to be the proximal interaction, ECM to be the intermediate interaction, and the secreting signaling molecule to be the distal interaction. Given that our approach is based on the neighbor search algorithm and that the input data consists solely of the spatial coordinates of the cells, considering the distal interaction will introduce error to the constructed CCI. Therefore, the initial CCI includes the proximal and intermediate-range interaction modes, and our algorithm combines two neighbor clustering methods, both of which have different purposes. The first is the hierarchical navigable small world (HNSW) algorithm [10], which is good at processing large numbers of data points and speeding up the construction of our spatial connectivity networks. The second is the Louvain algorithm [11], which can merge small clusters into larger ones, helping the search for intermediate-range cellular interactions. The implementations are summarized as follows:
Finding proximal interacting cells Kp.
Finding intermediate-range interacting cells Kir.
Integration of the initial CCI.
Figure 1. Demonstration of the seqFISH cell data features, where the blue overlay delineates the spatial coordinates of individual cells, the green overlay denotes the cell type classifications, and the red overlay represents the gene expression levels within each cell.
It is highly recommended that researchers combine Algorithm 1 with Figure 2 for a better understanding.
Algorithm 1: Construction of the cell adjacency matrix
Input: Cell spatial coordinates
Output: Cell adjacency matrix
Step 1: Find Kp proximal interacting cells.
1.1 Construct the spatial connectivity network.
1.2 kNNquery indexing Kp.
1.3 Form the proximal interaction adjacency matrix .
Adjp: [[N1, N2, …, Nc]1,[N1, N2, …, Nc]2,…[N1, N2, …, Nc](Nc) ]
Step 2: Find intermediate-range interacting cells.
2.1 Coalesce small clusters into a larger cluster.
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2.3 Form the inter-mediate range interaction adjacency matrix Adjir.
Step 3: Construct the cell adjacency matrix Adjc by combining Adjp, Adjir.
In Algorithm 1, in step 1.1, based on the input coordinates of the cells, a spatial connectivity network is constructed using the spatial proximity and geometric relationship. The result is shown in Figure 2A. The subsequent kNNquery in step 1.2 uses the spatial connectivity network to find Kp proximal interacting cells, defined as having potential direct cell–cell contact with the query cell. The green circle in Figure 2B is a demonstration of the randomly selected cell (red dot) and its interacting entities. The output of step 1 is the proximal interaction adjacency matrix Adjp∈R (Nc×Nc), where Nc is denoted as the number of input cells, the corresponding Kp index is 1, and the others are 0. Step 2 is the construction of the intermediate range interaction adjacency matrix. In step 2.1, the process involves grouping each cell with its neighbors and evaluating whether the maximum gain of modularity (∆Q) is greater than 0. If the gain is positive, the cell is assigned to the adjacent cell with the largest modular increment. Step 2.2 is the iteration of step 2.1, until ∆Q in Eq. (1) in Algorithm 1 no longer changes, being defined to be convergence. The result indicates that the cells within the same community are identified as potentially having intermediate-range interactions (ECM) with respect to the query cell, denoted as Kir. As a result, the number of potential interacting cells is Kp + Kir. The green circle in Figure 3C is the demonstration of a randomly picked cell (red dot) with its proximal interacting cells Kp and intermediate-range interacting cells Kir (green dots). In step 3, we can construct the adjacency matrix by
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In summary, Algorithm 1 distinguishes itself from the existing approaches relying on the kNN method, in which only is considered, resulting in the time complexity of O(Nc)2. Instead, taking both Kp and Kir into account, the potential ignorance of interactions that arise from various secretion patterns is prevented [12,13]. The overall time complexity is O(NclogNc + I ⋅(E+Nc )) where denotes the iterations in step 2.2 and denotes the number of edges constructed by the spatial connectivity network in step 1.1. Researchers can easily construct the cell level graph Gc by installing the packages "hnswlib" and "community" in Python.
Figure 2. Demonstration of the initial CCI construction using the seqFISH dataset. A. The purple dots represent the spatial locations of cells, and the spatial connectivity network generates the linkages between them. B. After step 1 in Algorithm 1, the randomly selected cell is in red in our demonstration, Kp (Kp = 2) returns in green and in the same size as the query cell does, denoted as the proximal interacting cells. (C) Cells in green inside the green circle (Kp+Kir) are defined to have potential interactions with the query cell in red, denoted as the proximal and intermediate-range interacting cells.
Figure 3. Construction process of Gg
2. Gene-level graph construction
The construction of the gene-level graph Gg is presented in Figure 3 and involves two main steps:
a. Construction of the cell-specific GRN:
Select Nselect (Nselect= Nlr+ Nrank) genes, comprising Nlr genes related to LR synthesis using standardized LR databases provided by Celltalkdb [14] and Nrank highly expressed genes.
Use the CeSpGRN backbone [15] to obtain the cell-specific gene interaction network, represented as cell-specific gene adjacency matrices (Nc graphs).
Apply Node2Vec [16] to generate features Fselect for the selected genes in each cell-specific graph.
b. Construction of the global GRN:
Combine the cell-specific graphs (NcjrG) into a global gene graph Gg.
Enrich the gene features by preserving the connections from the cell-specific graphs and using a meta-path-guided graph neural network [17,18] to derive new features F'select.
The final gene features Fg in Gg are obtained by integrating Fselect and F'select.
Researchers can use the following links to customize the gene level graph: https://github.com/PeterZZQ/CeSpGRN, https://github.com/eliorc/node2vec, and https://github.com/zhiqiangzhongddu/PM-HGNN.
B. Network architecture
Triple-enhancement-based graph neural network (TENET) is an end-to-end structure that could be utilized to retrieve ST data features as well as the graph structure to learn and reproduce. Gc and Gg are separately encoded in the graph convolution network (GCN) encoder in the first stage, obtaining the latent feature embeddings, denoted as Zc and Zg. Zg will traverse through the channel enhancement compensation block (CECB) and synergistic enhancement block (SEB). Before Zc moves on to the next stage, we integrate Zc and Zg to become Z(c+g). Then, Z(c+g) will traverse through CECB and the elective enhancement block (EEB). The whole process is illustrated in Figure 4.
Figure 4. TENET workflow. The GCN encoder extracts latent feature embeddings Zc and Zg from Gc and Gg. Before Zc is input into the subsequent blocks, we have an integration process converting Zc into Z(c+g). The subsequent triple-enhancement mechanism can be summarized as local-global-local enhancement. By doing so, the network can not only extract feature representations from the node itself but also from the graph structure. Before the decoding process, we fuse the latent feature embeddings from the enhanced block to have an accumulated effect. For the loss function, we employ the binary cross entropy (BCE) loss function for both cell and gene. Besides, due to the utilization of the triple-enhancement mechanism, if the model is to be subjected to noise or wrongly enhanced during the enhancement process, which is undesirable, it would struggle to converge. Consequently, we have the Loss"supervised" function that enables the model to combat noise and enhance its resilience effectively.
1. GCN encoder
The core of TENET's architecture is built upon the GCN [8], which serves as the foundation for extracting meaningful node representations from the input graph Gc and Gg and outputs the latent feature embeddings of cells and genes denoted as Zc and Zg, respectively. The calculation process of the two-layer GCN encoder is presented as
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2. Channel enhancement and compensation block (CECB)
The CECB in Algorithm 2 is a streamlined transition block; Z(c+g) and Zg are enhanced in the channel dimension, aiming to emphasize the relevant feature embeddings and mitigate the over-smoothing problem caused by the GCN encoder [19].
Algorithm 2. Channel enhancement and compensation block (CECB)
Input: Input feature: Z(c+g), Zg; Operation: Normalization: Batch normalization (BN), Activation function (ϕ,ω); Two fully connected layers [FC1, FC2].
Output: Z'(c+g), Z'g.
The results of the intermediate operations are represented using “temp” followed by different subscripts.
if Z(c+g) then
tempc+g= Z(c+g)-BN(Zc+g)
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Endif
Parallel operation for temp(c+g)1, temp(g)1.
temp(c+g)2 = ϕ(FC1(temp(c+g)1)), temp(g)2 = ω(FC2 (temp(g)1))
temp(c+g)3 = ϕ(FC1(temp(c+g)2)), temp(g)3 = ω(FC2(temp(g)2))
temp(c+g)4=temp(c+g)2*temp(c+g)3 , temp(g)4=temp(g)2*temp(g)3
if temp(c+g)4 then
Z'c+g = temp(c+g)1 + temp(c+g)4
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Endif
return Output: Z'(c+g), Z'g .
3. Synergistic enhancement block (SEB)
The enhancement of Z'g in the SEB is done jointly by the global branch enhancement and the local branch enhancement. In Algorithm 3, the global branch enhancement utilizes the graph attention network (GAT) [20], and the local branch enhancement acts as a buffer to integrate the feature embeddings extracted from the GAT.
Algorithm 3. Synergistic enhancement block (SEB)
Input: Input features:Z'g; Operation: Graph Attention Network (GAT), Convolution (Conv1), Batch normalization (BN), Activation function (ϕ,σ); Linear transformation (MLP, contains three fully connected layers [FC3,FC4,FC5]); Kernel list: K [1, 3, 5]; Group Size: G; Latent dimension: Lg1,Ll ,Lg2; Storage list: sl = []; Attention weight list:
awl [].
Output: Z''g .
The result of the intermediate operations are represented using “global” / “local” followed by different subscripts.
Global enhancement operation:
globalenhance1 = GAT(Z'g)[in channel (F'g),Lg1,attention heads]
The feature dimension turns into globalenhance1 ϵ RNg×(Lg1*attention heads).
Local enhancement operation:
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Global enhancement operation:
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4. Elective enhancement block (EEB)
The EEB in Algorithm 4 amplifies the latent feature embeddings by augmenting the spatial dimensions and leveraging adaptive feature selection [21], thereby bolstering the richness and caliber of the embeddings. The adaptive enhanced feature election mechanism in EEB bears a resemblance to the local branch in SEB.
Algorithm 3. Elective enhancement block (EEB)
Input: Input features: Z'(c+g); Operation: Adaptive average pooling(AAP), Convolution (Conv2,Conv3), Batch normalization (BN), Activation function (θ,ω); Two fully connected layers [FC6,FC7]); Latent dimension: .
Output: Z''(c+g)
The results of the intermediate operations are represented using “temp” followed by different subscripts.
temp1,temp2=Duplicate(Z'c+g)
Parallel operation for temp1, temp2.
temp3 = AAP(temp1), temp4 = AAP(temp2)
temp5=Concat(temp3, temp4).
Elective Enhancement operation: temp6= θ(BN(Conv2 (temp5)))
The feature dimension turns into temp6 ϵ RNC×L.
temp7, temp8=split(temp6)Parallel operation for temp7, temp8.
temp9= ω(FC6(Conv3 (temp7))), temp10= ω(FC6(Conv3(temp8))).
The feature dimension turns into temp10 ϵ RNC×F''c+g.
return Output: Z''c+g = FC7 (Z'c+g) * temp9*temp10.
5. Triple-enhancement fusionist decoder (TEFD)
Triple-enhancement fusionist decoder (TEFD) fuses the enhanced latent feature embeddings from the previous blocks as input to the inner product decoder [22]. The two decoders for cell and gene are parallel, denoted as and . The fusion mechanisms for the latent feature embeddings of cells and genes are:
Fusionc=sum(Zc+g, Z'c+g, Z''c+g). (4)
Fusiong=sum(Zg, Z'g, Z''c+g). (5)
The first terms in Eq. (4) and Eq. (5) Z(c+g) and Zg represent the raw latent feature embeddings obtained from the GCN encoder in subsection a. The second terms Z'(c+g) and Z'g undergo the CECB (primary enhancement) in subsection b. The third term Z''g is dual enhanced embeddings from SEB (secondary enhancement and tertiary enhancement) in subsection c. Analogously, Z''(c+g) is also a dual enhanced embedding when exported from EEB (secondary enhancement and tertiary enhancement) in subsection d. Each enhancement module incrementally amplifies inherent feature embeddings across different scales by progressively augmenting the channel dimensions, leading to a cumulative effect in Fusionc(g). To integrate the fusion features a step further, we have
Z'''c(g)= FC9(ReLU(FC8(Fusionc(g)))). (6)
Such a process discerns and extracts the salient features crucial for the subsequent decoding process.
The decoding process of Dc and Dg are summarized as follows:
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. The usage of the sigmoid activation is to limit the interaction scores between 0 and 1, representing the probability of the existence of an interaction between the corresponding cells (genes). Generally, TEFD ensures the interconnection and information flow among the blocks. The output of each block not only serves as input for the subsequent enhancement block but also contributes to the final reconstruction of CCI. By incorporating multi-scale information, a more nuanced understanding of cellular (genetic) interacting patterns can be achieved, allowing for a more accurate determination of communication establishment among cells.
6. Loss function
We employ the binary cross entropy (BCE) loss and propose a supervised loss in Algorithm 4. The BCE loss can achieve refinement in both the cell graph and gene graph and encourages the production of well-calibrated probabilities by penalizing large errors in the predicted probabilities [23].
To further correctly augment cell features, we have the supervised loss (Loss"supervised"). The inputs are the output of EEB and the output of the GCN encoder (after mapping and concatenating), the Z'(c+g) and Z(c+g), respectively, and its core concept is using the cosine similarity [24]. The embeddings at the same index are calculated in pairs, aiming to learn representations that bring similar cell embeddings in closer proximity [25] and dissimilar embeddings further apart. The triple-enhancement mechanism is employed to accentuate the specificity of the cells without significantly altering their representations. Therefore, a higher similarity value implies that the features are correctly enhanced and iteratively adjusted toward the desired enhancement, while a lower similarity value suggests the need for further adjustment to differentiate the features, leading to a modification of the cells' intrinsic properties. In addition to the embeddings being correctly enhanced, the triple-enhancement technique showcases remarkable robustness against noise. If noise is artificially simulated, the triple-enhancement structure can render the noise more prominent and distinguishable, and the cosine similarity between the enhanced and the original noise tends to be small. Therefore, TENET can effectively mitigate and suppress noise, thereby improving the reliability and stability of the model.
Algorithm 4. Supervised loss function
Input: Input features: Zc+g, Z''c+g.
Output: Loss value.
Emb=concat(Zc+g, Z''c+g)
Construct the similarity matrix →Sim(Emb, EmbT)
Elements of the same index → diagonal elements; Elements of different index → off-diagonal elements
return Output:
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Validation of protocol
Two hyper-parameter experiments A (train-test-split ratio) and B (tolerance to noisy data) are presented to validate the efficiency of TENET.
A. Train-test-split ratio
To validate the protocol, TENET’s model performance is compared with the two most related research DeepLinc and CLARIFY. For the validation, three ST datasets (seqFISH, MERFISH, and scMultiSim) are utilized, with a total of 120 iterations. Various train-test-split ratios (10%, 30%, 50%, 70%, and 90%) are employed to divide the data into training and testing subsets for evaluation. Figure 5A presents the segmentation results for a 70% train-test-split ratio (with a 30% proportion as the training set), where TENET is represented in red, CLARIFY in yellow, and DeepLinc in green. Figure 5B shows the AP results with the maximum and minimum values for each segmentation condition, with DeepLinc in green, CLARIFY in blue, and TENET in red. Notably, TENET consistently outperforms the other two models across all segmentation ratios on the three datasets.
Figure 5. Model performance comparison. A. Experiment conducted on three datasets under the segmentation of 70% (TENET in red, CLARIFY in yellow, and DeepLinc in green). B. The shadow in the shadow plot with the upper and lower boundaries extends to the maximum and minimum values of the AP results. DeepLinc in green, CLARIFY in blue, and TENET in red. Results are obtained from three repeated experiments.
Figure 6. Experiment conducted on three of the models using the dataset of seqFISH aiming to evaluate the model's tolerance to noise (Noise1 and Noise2). At each train-test-split ratio, the ratio of Noise1 and Noise2 gradually increases from 0 to 0.5 steps with a size of 0.1. The result in each subgraph is the average of three independent experiments under the condition of 120 epochs execution.
B. Tolerance to noisy data
The tolerance to noisy data experiment is designed to simulate poor data quality, including error-detection issues and missing or incorrectly detected edges in the experimental setup. In Noise1 experiment, a certain percentage of false edges are intentionally added to Gc and Gg. In Noise2 experiment, the existing edges are removed in proportion. We aim to mimic these scenarios and assess the model's ability to handle such challenges. These experiments are conducted under the condition of train-test-split ratios of 10%, 30%, 50%, 70%, and 90% on the dataset of seqFISH. The proportion of Noise1 and Noise2 increases from 0.0 to 0.5 with steps of 0.1, resulting in 36 AP values in each subplot in Figure 6. The values outside the bracket in Table 2 can represent the stability of the model with the increment of noise (Noise1 and Noise2) ratios. The value outside the bracket on one train-test-split ratio represents the elevation difference between Value1 (Noise1 and Noise2 = 0.0) and Value2 (Noise1 and Noise2 = 0.5). The value inside the bracket at one train-test-split ratio represents the average value of 36 AP, used to facilitate comparison among models.
Table 2. AP elevation difference of Noise1, Noise2 = 0.0, and (Noise1 and Noise2 = 0.5 on the same train-test-split ratio. The value in the bracket is the average AP from all proportions of the two noises on the same train-test-split ratio (36 AP values on one train-test-split ratio).
AP elevation difference
(Avg. AP)
Train-test-split ratio
10 30 50 70 90
Model
DeepLinc 30.22 (77.39) 26.91 (74.01) 23.74 (66.21) 22.09 (57.86) 17.27 (49.83)
CLARIFY 17.20 (83.86) 19.12 (81.04) 18.33 (75.13) 10.13 (69.22) 6.75 (61.64)
TENET 16.25 (88.89) 16.73 (85.37) 12.73(82.17) 12.12 (75.39) 7.02 (67.58)
* The best result for each model on each split ratio is highlighted in bold, and the second-best result is underlined.
General notes and troubleshooting
General notes
In Figure 7, there are two approaches to setting up the TENET environment. The simplest method is to utilize the provided environment.yml file (A); however, some dependencies may not be easily installed. Therefore, the recommended approach is the second way (B). Initially, create a Python virtual environment, then activate it and start installing the packages using the requirement.txt file. This file contains packages that can be downloaded without significant difficulty. Before installing the remaining packages, verify the CUDA version you are employing. If the CUDA version differs from this, the corresponding URL should be modified accordingly.
Figure 7. Environment preparation
To start training the model, use cd TENET to enter the program directory and use python main.py. The hyper-parameters can be adjusted according to the actual demand. The explanations of hyper-parameters in Figure 8 are as follows:
-m: "preprocess", this mode will construct the cell-level graph and gene-level graph using the provided method. This is necessary to generate the required graph structures before training the model. "train" this mode assumes that the cell-level graph and gene-level graph have already been constructed. It will use the provided graph structures to start the CCI reconstruction process.
-i: the input data frame path.
-o: the output results' path.
-t: train-test split ratio, the value ranges from 0 to 1. If default = 0.7, the test edges account for 70% percent of all.
-n, -k: these two hyper-parameters will be used when the mode (-m) is set to be "preprocess, where -n specifies the number of selected genes to be used when constructing the gene-level graph within each cell, -k controls the number of proximal interaction cells to be used.
-fp, -fn: the two noise ratios, their values range from 0 to 1. Introducing artificial noise to the constructed cell level graph can examine the model's tolerance to noises.
-lr: the value of the learning rate.
Figure 8. Hyper-parameters settings
Troubleshooting
The PyTorch version in TENET is 2.0.1 and the CUDA version is 11.7.
When installing PyTorch-related dependencies (torch-cluster, torch-scatter, torch-sparse, torch-spline-conv, and torch-geometric), they can be challenging to install and may require extended installation times. The recommendation is to download the respective .whl file via https://pytorch-geometric.com/whl (ensure you check the PyTorch version and CUDA version) and proceed with the installation.
Acknowledgments
This work is supported by the Guangdong Provincial Department of Education (2022KTSCX152), the Key Laboratory IRADS, Guangdong Province (2022B1212010006, R040000122), Guangdong Higher Education Upgrading Plan (2021-2025) with UIC Research Grant UICR0400025-21. The icons of the graphical abstract are created from https://BioRender.com.
Competing interests
The authors declare no conflicts of interest.
References
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Article Information
Publication history
Received: Jul 18, 2024
Accepted: Dec 25, 2024
Available online: Jan 21, 2025
Published: Feb 5, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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5,206 | https://bio-protocol.org/en/bpdetail?id=5206&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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Peer-reviewed
Cell-Sonar, an Easy and Low-cost Method to Track a Target Protein by Expression Changes of Specific Protein Markers
SB Sabrina Brockmöller
LM Lara Maria Molitor
FW Franz Worek
SR Simone Rothmiller
Published: Vol 15, Iss 3, Feb 5, 2025
DOI: 10.21769/BioProtoc.5206 Views: 44
Reviewed by: Alessandro DidonnaXin Xu Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Cells Aug 2024
Abstract
Different research methods aim to clarify the intracellular trafficking of target proteins or unknown pathways. Currently, existing methods are mostly complex and expensive, requiring expert knowledge. Detailed microscopy for protein co-localization detection or omic technologies, which provide holistic network data, are elaborate, mostly complex, and expensive to apply. Our protocol illustrates a method to track a target protein by detecting expression changes of user-selected marker proteins that directly or indirectly interact with the target. Modulation of protein expression indicates interactions between the target and marker protein. Even without co-localization analysis, the results of the protein expression change are the first insights into the target's fate. Moreover, the use of the cell-sonar is straightforward and affordable, and the results are rapidly available. Furthermore, this method could also be used to determine if and how pathways are affected by compounds added to the cells. In conclusion, our method is adaptable to a wide range of proteins, easy to apply, inexpensive, and expandable with substances that affect proteins.
Key features
• Easy-to-implement method to track intracellular proteins.
• Marker protein expression change demonstrates protein interaction.
• Combined data of all marker proteins is used to give an indirect overview of protein localization.
• This method is also applicable to different compounds and thus provides information about protein induction or influence on pathways.
Keywords: Protein expression changes Intracellular tracking Cell-sonar Pathway elucidation
Graphical overview
Workflow of cell-sonar
Background
Cell-sonar is an easy-to-implement method that allows indirect intracellular tracking of target proteins. The elucidation of secretory pathways has long been the primary objective of most research fields. Available methods for this are diverse microscopy techniques [1–4] or “omic” screenings [5]. These specific procedures result in high-definition data but, in almost all applications, they are complex, complicated, expensive, and elaborate. Our illustrated protocol is an alternative to these existing techniques, with the difference that cell-sonar is inexpensive, simple, and quick to apply. The results provide a first insight into protein localization.
The cell-sonar concept works universally for any application due to the principle of proteostasis, which regulates the protein balance in cells [6]. Several evolutionarily conserved signaling cascades for protein expression are controlled by proteostasis [6–8]. These protein networks are organized via signaling by recognition of stress responses and the maintenance of cellular proteins [7–9]. Internally, the endoplasmic reticulum (ER) controls the unfolded protein response (UPR) to each protein change [7] during its biosynthesis by regulating the ER capacity of nascent protein levels [7]. Signaling to the nucleus for the expression of required proteins is also orchestrated by UPR [10–12]. The signaling network located outside the ER is controlled by the mitogen-activated protein kinase (MAPK), and they interact in a synergistic manner [7]. The recognition of intracellular stress by MAPK-induced nucleus signaling is essential for gene expression [13]. This ability to dynamically detect changes, such as transgenic protein insertion or compound exposure, allows the detection of induced cellular protein expression. Exposure to a compound or expression of a transgenic target protein serves as an initiator of a stress response. Comparison of two cell lines that differ only in compound exposure or protein expression allows the detection of expression changes of interacting proteins, as shown in Figure 1. Altered intracellular protein levels provide a tracking indicator to observe secretory or other pathways of interest.
Figure 1. Schematic explanation of the cell-sonar principle. Mitogen-activated protein kinase (MAPK) and unfolded protein response (UPR) synergistically orchestrate signaling throughout the cell with subsequent induction of gene expression in the nucleus. (See details in Background section.) Highlighted in green are the components of the proteostasis network, which balances the synthesis, folding, and degradation of proteins to maintain the count of cellular proteins. MAPK and UPR are signaling networks that regulate any protein change that affects proteostasis processes. Highlighted in blue are the treatments that affect these balances. Chemical compounds or transgene proteins, which are not typically present in the cell, can result in an imbalance of the affected proteins as they interact (represented by blue dashed arrows). The proteostasis network regulates by altering gene expression to restore the balance. This is the reason why expression changes of interacting protein levels can be observed in direct comparison with untreated (green boxplot) and treated (blue boxplot) cellular proteins.
If there is a detectable expression change, it is the result of a protein interaction; if there is none, no protein interaction has happened. The results obtained from a few selected marker proteins provide a comprehensive overview of the fate of the target protein without the need to demonstrate the co-localization between a target and a marker protein. The advantages of cell-sonar are that it is adaptable to a wide range of proteins, easy to apply, and applicable to almost any cell line while it provides insights into the secretory pathway tracking in a short time.
The limitations include the fact that cell-sonar remains an overview. In order to make a reliable statement, it is crucial to use at least two marker proteins for one aspect and possess a detailed knowledge of the pathway and the proteins involved.
Materials and reagents
The following protocol instructions refer to the ER as the selected cellular area. Depending on the research question, the method can be expanded to additional cellular areas or various cellular pathways. In the following, we describe the basic principle and application of the method. We illustrate our protocol with two different cell lines, two different target proteins, and one compound to demonstrate the versatility of the cell-sonar method. Furthermore, we use the in-cell western method for the detection of protein expression changes but, of course, other methods such as on-cell western, western blot, or fluorescence-activated cell sorting analysis can also be used for this detection.
Biological materials
1. Chinese hamster ovary (CHO) cell line (Leibniz Institute DSMZ, ACC 110) and GH4C1 (Rattus norvegicus) cells (Genionics, Schlieren, Switzerland); both cell lines stably express different recombinant nicotinic acetylcholine receptors (nAChR). That means there are a) CHO cells, b) CHO cells expressing α12β1δϵ nAChR, [14], c) GH4C1 cells, and d) GH4C1 cells expressing α7 nAChR. Using identical cell lines for both host cells and genetically modified cells enables a direct comparison of protein levels
Reagents
1. MeOH (Merck/Sigma-Aldrich, catalog number: 1.06011.2500L)
2. Blocking buffer (LiCor, catalog number: 927-70001)
3. PBS pH 7.4, 1× dilution, without Mg and Ca (Gibco, catalog number: 10010-015)
4. Tween 20 (Merck/Sigma-Aldrich, catalog number: P1379-100mL)
5. Cell tag stain 700 (LiCor, catalog number: 926-41090)
6. Nicotine (Merck/Sigma-Aldrich, catalog number: N3876-25mL)
7. Primary antibodies: calnexin (CN) (Abcam, catalog number: ab133615), BiP (Abcam, catalog number: ab213258), Sil1 (Abcam, catalog number: ab228868), Hrd1 (Abcam, catalog number: ab249578), UGGT1 (Abcam, catalog number: ab124879), and GAPDH (Abcam, catalog number: ab8245)
8. Secondary antibodies: IRDye 800 rabbit (LiCor, catalog number: 92632211) and IRDye 800 mouse (LiCor, catalog number: 2632210)
9. Culture medium F-12 Nut Mix + GlutaMaxTM (Gibco, catalog number: 31765-027)
10. Fetal calf serum (FCS) (Gibco, catalog number: 10270-106)
Solutions
1. PBST (0.1% Tween-20 in PBS)
2. Primary antibody dilution 1:200 in blocking buffer
3. Secondary antibody dilution 1:800 in blocking buffer
4. Nicotine cell treatment (see Recipes)
Recipes
1. Nicotine cell treatment
Reagent Final concentration Exposure time
Nicotine 30 μM 24 h
Laboratory supplies
1. Black 96-well microplates with clear bottoms (Greiner Bio one, catalog number: 655986)
Equipment
Odyssey CLx Imager (LiCor, catalog number: 9140)
Note: Odyssey CLx is no longer available, the successor model is Odyssey DLx Imager, catalog number: 9142.
2. KS 260 control IKA (Sigma-Aldrich, catalog number: Z341835)
3. Fume hood HERA Safe KSP (Thermo Scientific, catalog number: 17168075)
Software and datasets
1. Image Studio software (version 5.2 LiCor)
2. Prism [version 9.5.1 (733) GraphPad]
3. Excel (version 16.0.10415.20025 Microsoft Excel 2019 MSO)
4. BioRender (https://www.biorender.com/). The following figures were created using BioRender: Graphical overview, BioRender.com/s04v667; Figure 1, BioRender.com/w21c279; Figure 2, BioRender.com/i62d658
Procedure
A. Determine the linear regression for cell count normalization
1. Seed CHO cells in culture medium and 16.5% FCS, and GH4C1 cells in the same medium with 10% FCS supplement in different black 96-well microplates in linearly increasing cell counts: 0, 5,000, 10,000, 15,000, 20,000, 25,000, 30,000, 35,000, 40,000, 45,000, and 50,000 cells. Apply biological and technical triplicates in corresponding culture media each with 100 μL as the final volume per well.
2. Incubate plates for 24 h at 37 °C, 5% CO2, and 90% humidity.
3. Perform cellular fixation and permeabilization in a fume hood with 50 μL of 100% MeOH (-20 °C) per well for 10 min at room temperature.
4. Gently wash each well three times with 150 μL of PBS for 5 min.
5. Block every well with 150 μL of blocking buffer for 1.5 h at room temperature at 180 rpm.
6. Apply cell tag stain 700 solution, necessary for normalization on cell counts, at 50 μL/well and incubate for 1 h at room temperature at 180 rpm, protected from light. Therefore, dissolve cell tag stain 700 in 100 μL of PBS and incubate for 30 min at room temperature protected from light; the concentration per well is 0.2 μM.
7. Gently wash each well three times with 150 μL of PBST for 5 min.
8. Add 100 μL of PBS per well before scanning plates with Odyssey CLx Imager using the Image Studio software.
9. Scan the plate with the following settings: resolution = 169 μm, quality = medium, and focus offset = 3.8 mm.
Note: Focus offset needs to be adjusted for different plates.
10. Match each scanned fluorescence per well utilizing the special grid of the Analysis tab of the software menu.
11. Export the data as an Excel report and use the detected fluorescence signals to determine the linear regression for cell count normalization.
B. Perform in-cell western for marker protein expression level detection
1. Seed 30,000 CHO and GH4C1 cells in different black 96-well microplates in biological triplicates and four technical replicates in corresponding culture media each with 100 μL as the final volume per well. In the case of nicotine treatment, the final concentration of nicotine in the wells was 30 μM.
Note: Nicotine treatment was performed in our case regarding its anticipated effect on receptors and as an example of compound application in the cell-sonar method.
2. Incubate plates for 24 h at 37 °C, 5% CO2, and 90% humidity.
3. Perform cellular fixation and permeabilization with 50 μL of 100% MeOH (-20 °C) per well for 10 min at room temperature.
4. Gently wash each well three times with 150 μL of PBS for 5 min.
5. Block every well with 150 μL of blocking buffer for 1.5 h at room temperature at 180 rpm.
6. Apply 50 μL of primary antibody solution/well and incubate for 2.5 h at room temperature at 180 rpm. Perform each microplate with controls, which are shown in Figure 2.
7. Gently wash each well three times with 150 μL of PBST for 5 min.
8. Apply cell tag stain 700 solution, necessary for normalization on cell counts, at 0.2 μM concentration and secondary antibody dilution at 50 μL/well and incubate for 1 h at room temperature and 180 rpm protected from light. Do not add cell tag stain 700 solution to the control wells of the secondary antibody signal alone (see Figure 2).
Figure 2. Schematic microplate overview with controls and samples. The plate layout shows how to handle controls (I–IV) and samples (V–X). I. Add three wells with blocking buffer only. II. Add three wells with cells in blocking buffer only. III. Add two wells (in sum 12 wells) for each solution of all individual primary antibodies on cells in blocking buffer. IV. Add three wells (in sum 6 wells) for each solution of all individual secondary antibodies on cells in blocking buffer all without adding cell tag stain 700. V–X. Performed for marker protein expression level detection; each marker protein in a biological triplicate (n = 3) and with four technical repeats, for a total of 12 wells. To these wells, primary antibody solution, secondary antibody solution, and cell tag stain 700 were added. This microplate had to be performed eight times in total for host CHO and GH4C1 cells, for host CHO and GH4C1 cells under nicotine treatment, for transduced CHO and GH4C1 cells, and for transduced CHO and GH4C1 cells under nicotine treatment.
9. Gently wash each well three times with 150 μL of PBST for 5 min.
10. Add 100 μL of PBS/well before scanning plates with Odyssey CLx Imager using Image Studio software.
11. Scan the plate with the following settings: resolution = 169 μm, quality = medium, and focus offset = 3.8 mm.
Note: Focus offset needs to be adjusted for different plates.
12. Match each scanned fluorescence per well by utilizing the special grid of the Analysis tab of the software menu.
13. Export data as an Excel report and utilize the detected fluorescence signals for analysis of the expressed protein levels.
Data analysis
Raw data may be analyzed in GraphPad Prism, as in our case. As visualized in Figure 2, multiple controls are mandatory for data analysis. Controls I–III of Figure 2 should not detect signals stronger than the sample fluorescence values. Control IV of Figure 2 shows the background signals required to average the secondary rabbit and secondary mouse IRDye 800 intensities. We recommend a minimum of three technical replicates per control. For data analysis, initially, the mean of the mouse or rabbit background was subtracted from all acquired data. To normalize the raw data to an accurate cell count, the user is required to perform a linear regression, as described in section A. In our example, the fluorescence signals were normalized to a cell count of 30,000 cells per well.
The preprocessed data are then used to generate boxplots. Each marker protein is represented by a boxplot of 12 data points. Boxplots were presented for host cells with transduced cells without treatment and both cells under nicotine treatment. Additionally, a two-sample t-test with dependent samples was conducted. p-values below 0.05 were considered statistically significant.
Any case of significant expression changes demonstrates a protein interaction between the marker protein and the target protein or the influence of the applied compound. The results are compared to the current literature to infer ATP-dependent or calnexin cycle–dependent associations. The sum of the data enables the tracking of proteins within the ER and throughout the cell, in the case of additional regions.
Validation of protocol
This protocol has been used and validated in the following research article:
• Brockmöller et al. [15]. Cell-sonar, a Novel Method for Intracellular Tracking of Secretory Pathways. Cells 13(17), (complete publication, three chosen cellular areas: the ER, between the ER and Golgi, and the endocytic pathway, with CHO cells and transgene α12β1δϵ nAChR).
Additionally, this protocol has been used and validated in previously unpublished experiments with GH4C1 cells and the transgene α7 nAChR. Initially, we performed a western blot from purified protein solutions to demonstrate that α7 nAChR is expressed in the transgene GH4C1 cell line but not in the GH4C1 host cell line (Figure 3). The cell-sonar was performed with UGGT1, CN, Hrd1, BiP, Sil1, and GAPDH, which are marker proteins for the ER area [15], in the presence and absence of nicotine treatment. The results are presented in Figure 4. As described in the Data Analysis section, changes in protein expression were detected for five marker proteins, which means that the presence of α7 nAChR can be indirectly detected without western blot analysis. Furthermore, the ER area is divided into ATP-dependent (Figure 4A) and calnexin cycle–dependent domains (Figure 4B). The receptor also interacts differently with these marker proteins without and under nicotine treatment. Our protocol has also been applied to our laboratory procedures to gain new insights into some secretory pathways under different aspects and target proteins.
Figure 3. Western blot of the α7 subunit of nAChR from transgene (+) and host GH4C1 (-) cells. Western blot was performed from 57 ng of purified protein solutions of both GH4C1 cell lines. M shows the marker, 50 and 70 kDa bands are indicated.
Figure 4. Expression changes of in-cell western for BiP, Sil1, GAPDH, UGGT1, CN, and Hrd1 as endoplasmic reticulum (ER) markers. Boxplots of normalized fluorescence signals of A) BiP, Sil1, and GAPDH as markers for ATP-dependent associations show expression changes between α7 nAChR-expressing GH4C1 cells and untransduced controls in the presence or absence of nicotine, whereas GAPDH was not increased under either condition. B) UGGT1, CN, and Hrd1 as markers for calnexin-cycle dependent associations show different expression levels, whereas CN was not significantly increased under nicotine treatment. Data are combined from biological triplicates with four technical replicates each, *** means significance with a p-value below 0.001.
General notes and troubleshooting
General notes
1. Each aspect that should be investigated requires at least two marker proteins. The result from one marker may be a coincidence and is thus not sufficient. Thus, we recommend a minimum of two marker proteins for a particular aspect.
2. It is important to allow enough time to find suitable marker proteins. Without knowledge of secretory pathways or intracellular understanding, conclusions could be incorrect. The application of the cell-sonar in the laboratory is straightforward; however, the theoretical foundation requires thorough literature research.
3. Interacting proteins do not require direct contact to function, and communication is possible through signaling throughout the cell. Cell-sonar demonstrates the interaction of proteins but not the co-localization of those.
Troubleshooting
Problem 1: Clustered proliferation of some cell lines, like HEK cells. If cell lines proliferate in a very clustered manner, whole cell clusters may break from the well bottom at the wash steps and distort the following results.
Possible cause: The number of plated cells is too high and/or the washing steps are too excessive.
Solution: Plate a lower cell count of HEK cells and execute each washing step with the utmost care.
Acknowledgments
We would like to thank Prof. Dr. Horst Thiermann for providing the laboratories and working equipment. Further, we would like to thank Dr. Hildegard Mack for her extensive support.
Funding: The authors declare that no funds, grants, or other support were received during the preparation of this manuscript.
Previous work or the original research paper in which this protocol was described and validated: Brockmöller, S.; Seeger, T.; Worek, F.; Rothmiller, S. (2024). Cell-Sonar, a Novel Method for Intracellular Tracking of Secretory Pathways. Cells, 13(17): 1449, doi:10.3390/cells13171449 [15].
Competing interests
The authors have no relevant financial or non-financial interests to disclose.
References
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Ghisaidoobe, A. and Chung, S. (2014). Intrinsic Tryptophan Fluorescence in the Detection and Analysis of Proteins: A Focus on Förster Resonance Energy Transfer Techniques. Int J Mol Sci. 15(12): 22518–22538.
Axelrod, D. (2008). Chapter 7 Total Internal Reflection Fluorescence Microscopy. Methods Cell Biol. 89: 169–221.
Herzenberg, L. A., Parks, D., Sahaf, B., Perez, O., Roederer, M. and Herzenberg, L. A. (2002). The History and Future of the Fluorescence Activated Cell Sorter and Flow Cytometry: A View from Stanford. Clin Chem. 48(10): 1819–1827.
Horgan, R. P. and Kenny, L. C. (2011). ‘Omic’ technologies: genomics, transcriptomics, proteomics and metabolomics. Obstetr Gynaecol. 13(3): 189–195.
Balch, W. E., Morimoto, R. I., Dillin, A. and Kelly, J. W. (2008). Adapting Proteostasis for Disease Intervention. Science. 319(5865): 916–919.
Hotamisligil, G. S. and Davis, R. J. (2016). Cell Signaling and Stress Responses. Cold Spring Harbor Perspect Biol. 8(10): a006072.
Hipp, M. S., Kasturi, P. and Hartl, F. U. (2019). The proteostasis network and its decline in ageing. Nat Rev Mol Cell Biol. 20(7): 421–435.
Meiners, S. and Ballweg, K. (2014). Proteostasis in pediatric pulmonary pathology. Mol Cell Pediatr. 1(1): e1186/s40348–014–0011–1.
Shamu, C. E. and Walter, P. (1996). Oligomerization and phosphorylation of the Ire1p kinase during intracellular signaling from the endoplasmic reticulum to the nucleus. EMBO J. 15(12): 3028–3039.
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Peters, A., Nawrot, T. S. and Baccarelli, A. A. (2021). Hallmarks of environmental insults. Cell. 184(6): 1455–1468.
Morrison, D. K. (2012). MAP kinase pathways. Cold Spring Harb Perspect Biol. 4(11): a011254.
Brockmöller, S., Seeger, T., Worek, F. and Rothmiller, S. (2023). Recombinant cellular model system for human muscle-type nicotinic acetylcholine receptor α12β1δε. Cell Stress Chaperon. 28(6): 1013–1025.
Brockmöller, S., Seeger, T., Worek, F. and Rothmiller, S. (2024). Cell-Sonar, a Novel Method for Intracellular Tracking of Secretory Pathways. Cells. 13(17): 1449.
Article Information
Publication history
Received: Oct 15, 2024
Accepted: Dec 19, 2024
Available online: Jan 14, 2025
Published: Feb 5, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Biochemistry > Protein > Interaction
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5,207 | https://bio-protocol.org/en/bpdetail?id=5207&type=0 | # Bio-Protocol Content
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A Micro-Computed Tomography-Based Simplified Approach to Measure Body Composition, Osteoporosis, and Lung Fibrosis in Mice
ML Madeleine B. Landau *
BZ Binghao Zou *
ZY Ziqi Yang
BR Brian G. Rowan
MA Muralidharan Anbalagan
(*contributed equally to this work)
In Press, Available online: Jan 19, 2025
DOI: 10.21769/BioProtoc.5207 Views: 40
Reviewed by: Komuraiah MyakalaKanchan Bhasin Anonymous reviewer(s)
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Abstract
Micro-computed tomography (micro-CT) is a powerful, non-destructive imaging technique that creates high-resolution 3D images of the internal structures of small animal models such as mice and rats. Familiarizing oneself with micro-CT imaging and data analysis can be overwhelming without easy-to-follow, clear instructions. Training on new instruments is often a task exclusive to a select subset of researchers, leaving the majority of potential trainees without a technical grasp of how to navigate the instructions. This protocol on the use of micro-CT aims to bridge that gap by providing a clear, step-by-step guide to acquire and analyze micro-CT images from mice for quantitative data. By exclusively detailing the necessary procedural steps from start to finish and overcoming complex user interfaces during imaging operations and analysis, this protocol will equip new micro-CT users with the ability to measure mouse body composition (bone, body fat, and lean muscle mass) and identify and quantify lung fibrosis. This approach applies to researchers with a basic understanding of medical imaging, animal care, and software analysis.
Key features
• Analysis of tissue-specific body composition using mice as model organisms.
• An easy-to-follow guide for novice users of high-resolution micro-computed tomography imaging systems.
• Enhances accessibility, workflow, standardization, training, and breadth of application in the research community.
• Effectively employing non-invasive live imaging allows for a longitudinal study of tissue architecture for examining age-related changes in vivo.
Keywords: Micro-computed tomography Micro-CT Murine models Body composition Bone mineral density Adipose Lean tissue Lung fibrosis Tissue structure
Graphical overview
Background
The role of small animal models in basic science research has rapidly expanded amidst the emergence of novel quantification techniques [1]. Precision measurement in such models has endeavored to unearth underlying mechanisms of action in estrogen-receptor signaling and innovative modes of drug delivery for quality improvement [2,3]. Genetically modified mice have an indispensable purpose in assessing the role of specific genes in key physiological functions and phenotypic changes that are often measured through histological analysis [4,5]. Nonetheless, such models may fail to provide either pertinent longitudinal data, an important consideration for examining structural development (fat, muscle, bone), or pathological changes in tissues such as lung fibrosis.
As an alternative approach to examining in vivo features at a fixed point in time, non-invasive techniques that allow for periodic imaging over time can quantitatively measure varied physiologic and pathologic phenotypes [5]. The benefits of live imaging of murine structures include quantifiable measurements for organic tissue-dependent changes at specific growth stages without inflicting harm or interruption to the life cycle. Non-invasive tools also enhance experimental designs by yielding a series of data points over the entire study duration, limiting the overall cohort size needed, and allowing more precise measures of inter-animal variability [6].
One of the most frequently utilized tools for diagnostic imaging is X-ray computed tomography (CT), which relies on employing X-ray beams in direct contact with the organism to create a cross-sectional tomographic representation. Hounsfield units (HU) is a relative quantitative measurement of radio density used in the interpretation of computer tomography images in the linear scale; the radio density of distilled water at a standard temperature and pressure is defined as 0 HU, whereas the radio density of air is -1,000 HU. A phantom with a known radio density calibrates machines, ensuring they provide the correct HU. Most animal tissues do not exceed +2,000 HU and standard clinical limits range from -1,000 to +3,071 [7].
In pre-clinical research, the ability of the micro-CT scanner to elicit information on body composition comes highly appreciated, enhancing surveillance of disease development, classification, and ramifications [8]. Whether analyzing bone mineral density for studying osteoporosis, distinguishing airspace changes for characterizing lung fibrosis, or comparing adipose volume change, non-invasive in vivo imaging using micro-CT scanners enables high spatial resolution with longitudinal monitoring [9].
Without a standardized protocol for utilization, not only does the reproducibility of research suffer, but it also hinders the ability to build upon the ever-evolving field of structural analysis and to develop an eventual gold standard for body composition characterization. Simple method articles serve as valuable resources for researchers who wish to incorporate micro-CT into their studies.
In this article, we propose an easy-to-follow guide for micro-CT users to reconstruct the physical structure of mice to obtain standard non-invasive imaging in mice-based long-term experiments. Our protocol improves upon existing methods by providing accessible, step-by-step instructions that minimize the need for specialized training. It simplifies workflows into manageable tasks, promotes standardization with many visual aids and troubleshooting guidance, and improves reliability. This protocol details a reproducible manner by which researchers can precisely attain data on lean tissue, adipose, bone structure, bone mineral density, and airspace quantification in the lung, enabling the investigation of gene-based translational applications.
Materials and reagents
Biological materials
1. C57BL/6 mice, 3 months old (C57BL/6J from The Jackson Laboratory, Bar Harbor, ME)
Reagents
1. FlurisoTM, isoflurane (VETone, catalog number: 502017), store at 20–25 °C for up to five years
2. Oxygen [50 pounds per square inch gauge (psig) (345 kPA) to 60 psig (414 kPA) required] (air gas)
Note: Oxygen can be a fire hazard, so always turn it off when not in use. Do not expose oxygen cylinders to temperatures higher than 50 °C. The oxygen tank should be securely fastened upright to prevent falling off and kept in a well-ventilated area completely separated from flammable materials and heating sources. Always keep the oxygen tank valve protected with the cap when not in use and ensure proper labeling indicating the contents as OXYGEN and whether it is full or empty. Never attempt to handle a damaged or leaking cylinder. Be aware of your laboratory-specific safety procedures while handling the oxygen tank.
3. Bleomycin solution 10 mg/mL in water (Sigma, catalog number: B7216); should be protected from light and stored at -20 °C. Mice were given 60 µL of bleomycin (1.25 U·kg-1 body weight) via oropharyngeal route to induce a fibrotic response that is more pronounced at day 21 after the treatment [10]. The Institutional Animal Care Use Committee (IACUC) approved this bleomycin-induced lung fibrosis protocol (Protocol ID: 1874)
Equipment
1. RAS-4 anesthesia system (PerkinElmer, catalog number: CLS146737)
2. Quantum GX2 micro-CT (PerkinElmer, catalog number: CLS149276)
3. PC: DELL Precision 5820 tower XCTO high-performance acquisition computer, Windows®10 Pro, Intel® (R) Xeon (R) W-2123 processor, NVIDIA Quadro P2000 5GB graphics card, 32 GB 2666 MHz RAM, 8 TB HD.
Note: Researchers working with animals in our vivarium must have taken proper animal training and lectures. They must wear appropriate personal protective equipment (PPE), including a dedicated lab coat or disposable gown, gloves, mask, and head and shoe covers. All PPE is available at the entrances to our animal facilities. Additionally, researchers must take radiation safety training before operating the micro-CT instrument. We recommend that researchers follow the institution-dependent regulations and use PPE accordingly.
Software and datasets
1. Quantum GX2 Image Analysis Software (PerkinElmer)
2. PerkinElmer Database, version 3.5.3.110 (free software with the purchase of Quantum GX2)
3. Analyze 14.0 (AnalyzeDirect, Inc., Lexana, KS, USA, license needed, https://analyzedirect.com/analyze14/)
4. Prism v9.3 (GraphPad, 11/15/2021, license needed, https://www.graphpad.com/). To perform a t-test in Prism: Launch the software and create a new project. Choose the Column table format and enter the data into the appropriate columns. Click the Analyze button and, under the Column Analyses group, select t-tests (and nonparametric tests).
5. BioRender (https://www.biorender.com/). The following figures were created using BioRender: Graphical overview, Created in BioRender. Landau, M. (2024) https://BioRender.com/c08f795.
Procedure
A. Micro-CT calibration and scanning for body composition
1. Turn on the Quantum GX2 Micro-CT imaging system and switch the safety key to On (Figure 1A).
Figure 1. Quantum GX2 micro-CT imaging system. A) Quantum GX2 micro-CT system with the computer in our facility. B) RAS-4 rodent anesthesia system. C) Anesthetized C57BL/6 mouse placed on the mouse bed with the nose inserted inside the nose cone continuously receiving anesthesia from the RAS-4 anesthesia system. D) Mouse’s whole body scan with coronal, sagittal, and axial views.
2. Open the Quantum GX2 Image Analysis Software and click the Warm-Up button in the lower left corner.
Pause point: The warm-up process takes 15 min if the duration of imaging system disuse is <1 month. For durations of 1–3 months, it takes 40 min; for >3 months, it takes 2 h.
3. Critical: In the database window, click Create New Database to set the folder for saving the imaging data files. Then click Create New Sample and enter the information of the new sample to set the folder for saving the data of the particular scan. Click the sample that will be scanned next and click Set Image Save Location.
4. The Quantum GX2 micro-CT may require gain or HU calibration if the indicator turns yellow or red before animal scanning.
a. To complete the gain calibration, select Gain Calibration from Options of the control panel. Select the required Scan mode, Filter, and Voltage combination that is needed to be calibrated and click Execute (Figure S1A).
Note: The indicator should turn green after the gain calibration process is complete.
b. Set up a HU calibration water phantom by adding 25 mL of DI water into a 50 mL conical centrifuge tube. Place the water phantom into the Quantum GX2 micro-CT and configure the control panel to the required Scan mode, Filter, and Voltage combination. Create a new sample to store the scan file. Enter live mode by clicking the Live Mode button. Stage control on the front panel is used to adjust the position of the bed, and rotation control is used to confirm that the conical centrifuge tube is centered in the field. Click Start Scan to complete a scan.
c. After scanning, right-click the sample in the series information and select Launch HU Calibration. Select Settings and VOX Number Calibration (V) to launch an ROI selection window. Move the green square that represents AIR to the air area in the conical centrifuge tube and the TARGET green square to the water area. In the CT number adjustment, click ROI read for AIR CT and TARGET CT and click OK (Figure S1B). Close the measurement window. On the control panel, select Options and HU Calibration Settings. Find the scan mode that requires HU Calibration, choose the most recent calibration file, and the indicator should turn green (Figure S1C).
5. Before beginning animal scanning, anesthetize the mouse with 2.5% isoflurane in the RAS-4 rodent anesthesia system induction chamber.
Note: It takes 2 min to fully anesthetize a mouse (Figure 1B).
6. Place the anesthetized mouse on the mouse bed and close the imaging system door (Figure 1C).
Critical: Place only one mouse into the mouse bed at a time. Individual micro-CT scanning is encouraged for animal safety and best imaging results. The mouse's nose must be fully inserted into the nose cone to ensure consistent anesthesia delivery through inhalation of isoflurane (Figure S2).
7. Adjust the scan settings in the control window as follows:
a. For bone marrow density (BMD): voltage: 90 kV; current: 88 µA; acquisition: 36; recon: 36; scan mode: standard, 18 s; display settings: off; X-ray filter: Cu 0.06+Al 0.5.
b. For body composition (other): voltage: 45 kV; current: 133 µA; acquisition: 36; recon: 36; scan mode: standard, 18 s; display settings: off; X-ray filter: none.
8. The field of view (FOV) will present the size of each micro-CT scan. A FOV of 36 mm was used, which requires stitching five separate scans together to capture the entire mouse.
Note: A larger FOV of 72 mm requires just three scan stitches to obtain the entire coverage. However, increasing from 36 to 72 mm for the FOV will move the camera further away from the mouse, subsequently reducing the resolution of the image. For larger rodents, such as rats or rabbits, a larger FOV may be the only plausible option.
9. Each of the four changeable filters can be used for optimal imaging of different structures:
a. No filter (open): low contrast samples at low voltages.
b. AI 0.5 mm: Low contrast samples.
c. AI 1.0 mm: Soft tissue (fat analysis).
d. AI 0.5 mm + Cu 0.06 mm: Standard CT scanning.
e. Cu 0.1 mm: Dense samples at high voltages.
10. Set the body orientation as desired. Click Initialize in the control window to initialize the stage control.
11. At the bottom of the control window, click Live Mode to turn on the live mode and use the stage controls on the front control panel to locate the animal in the center of the live window. Use the “rotation control” to ensure the animal is centered. Move the stage to position the nose of the mouse to the center of the FOV and record the stage location.
Note: This should be the starting point of the five stitches.
12. To set a stitching job, select Options > Job Scan Settings and click New in the Job Menu (Figure S3). Press Add in the Row Menu to create a 5-Job scan. Enter the recorded number of the position of the nose as the starting point in the stage Z(mm).
Pause Point: The Quantum GX2 can stitch up to five scans, which is sufficient to reconstruct the micro-CT image for the body of an adult mouse at a 36 mm FOV.
Pause Point: The range of the stage for stitching is 2–206, so it may be necessary to adjust the location of the nose cone to ensure that the starting and the end of the five stitches do not exceed this range.
13. Change the stitching to On and adjust the other parameters that are required for the scan. Click OK.
14. In Scan Acquisition settings, select the job just created and confirm the parameters are correct for the scan.
15. Start scanning by clicking the CT Scan button. When the machine status becomes stand by, remove the animal and ensure the X-RAY ON light is off.
Note: Scans are viewable from the coronal, sagittal, and axial viewpoints (Figure 1D).
16. A phantom plot is required to calculate the standard curve for BMD quantification. A phantom has five different densities (0, 50, 200, 800, and 1,200 mg HA/cc) of hydroxyapatite (HA). Use the same setting for the phantom as for BMD scanning (Figure S4A).
B. Micro-CT phantom plot generation
1. To analyze 3D scans generated from the Quantum GX2 micro-CT, the associated software, the Quantum GX2 Database application, must be opened on the desktop. The user must consent to allow an existing database to connect. Once the data via USB is transferred to the computer, and upon enabling the existing database to connect to the Quantum GX2 micro-CT program, each scan becomes viewable.
Pause Point: If Analyze 14.0 is not installed on the same computer used to control Quantum GX2 scanning, the data must be transferred to a computer with Analyze 14.0 software. To transfer data, copy the entire database folder (the same directory as shown on the database location) or select Export File to export the selected sample.
2. In the Quantum GX2 Database application, select a given image file for loading, then double-click to open and enlarge the image. The unique file name should have a signature sample ID, sample name, date of birth, sex, and weight.
3. Open Analyze 14.0 on the Desktop and click New Workspace. Select the Input/Output function on the right-hand side. The user must manually search for the corresponding file opened previously in the Quantum GX2 Database based on the file ID number. The .VOX file should be chosen for file type. Then, select Calculate it.
4. Once the image appears in the workspace, click Load Volume in the bottom left-hand corner of the screen. Click Exit, then Rename. The file desired for analysis may then be designated according to the preferred naming system.
5. Select Transform on the right-hand side to localize the region of interest within the image. Choose Spatial Transforms and Subregion function within, then click Extract Sub-Volume to delete unnecessary regions and save computing power. Click Save Volume in the lower left-hand corner of the screen, which saves the transformed image in the new workspace.
6. Select Process from the right-hand side of the screen. Click Spatial Filter for the Process Type, Median for the Filter Type, Kernel Size 3 for the Subtype, and Automatic for the Preview Type. Then, click Process Volume to generate a smoother image and reduce background noise, overall increasing image quality. Press Save Volume.
Pause Point: At this point, three images are saved in the workspace: the original image from the hard drive import, the image after applying the Transform function, and the image after applying the Process function.
7. Select Segment on the right-hand side. With the five standards in view, select Fabricate Shapes under Manual. For dimensions, click 3-D, and for Shape, click Cylinder in the Object to Create tab, then check the box adjacent to Filled.
8. Adjust the radius of the cylinder within the standard to accurately sample the density of the standard. Adjustments of X, Y, and Z coordinates can be made through settings in the Location and Size section. When the desired size and location adjustments are complete, select Apply to fix the volume and placement of the cylinder.
9. Click Add Object and drag the new Object from the original segment to a new standard. Once in position, click Apply. Repeat this step so all standards contain a cylinder (Figure S4B, C).
10. Choose Save Object and open Measure on the right-hand side.
11. All five circular density samples should be visible within the confines of each standard. Select 3D under the Sample Type, then choose Enabled Objects. Under the Stats to View section, click Size Intensity and pick Mean. Check the box for auto log stats, and under Sample Options, click Sample Enabled Objects. The mean value is generated from all five standards, visible below the scanned image.
12. Open a new Excel file spreadsheet and copy the computed values into the Excel file. Title one column Object Name and input the standard densities in this row. Create an adjacent column for Mean and then plot the standard curve with the standards on the X-axis and means on the Y-axis. Use Excel to calculate the equation of the standard curve. The trendline of this graph can be referenced when prompted for scaling parameters (“SigmaCT” = slope, “BetaCT” = offset) (Figure S4D).
C. Quantification of bone mineral density (BMD) of mouse femur
Upload the mouse data and start the software applications in the same manner as outlined in section B.
1. Select an image file for loading, then double-click to open and enlarge the image. All captured images should have a unique file name including a signature sample ID, sample name, date of birth, sex, and weight.
2. Open Analyze 14.0 on the Desktop and click New Workspace. Select Input on the right-hand side. Manually search for the identical sample ID number.
3. Find the corresponding file opened previously in the Quantum GX2 Database based on the file ID number. The .VOX file should be selected for file type. Then, click Calculate it.
4. Once the image appears in the designated workspace, click Load Volume in the bottom left-hand corner of the screen. Click Exit, then Rename (if desired).
5. Select Transform on the right-hand side to localize the region of interest within the image. Choose Spatial Transforms and the Subregion function within, then click Extract Sub-Volume to compute the new volume. Click Save Volume in the lower left-hand corner of the screen.
6. Select Process from the right-hand side of the screen. Click Spatial Filter for the “Process Type,” Median for the “Filter Type,” Kernel Size 3 for the “Subtype,” and Automatic for the “Preview Type.” Then, click Process Volume. Press Save Volume.
7. Select Segment on the right-hand side. Under the Semi-Automatic tab, click Threshold Volume and adjust the threshold to find the best range for the object density imaged. Once the optimal threshold range is attained, choose Threshold Object to isolate the bones (Figure 2A).
Figure 2. Segmentation of the mouse bone (Femur) and bone mineral density (BMD) quantification. A) Image shows the threshold to segment the mouse skeleton, B) Image showing that the manual trace tool was used to select the femur (green), C) Image shows the removal of other bones with only the femur present, D) This image shows the use BMA tool to segment cortical and trabecular bone E) This image shows the window for calculating and analyzing the density of the femur.
8. Select Manual, which enables manual tracing of the desired Object (bone). Press Add Object (green), and then trace around excess structures outside the object of interest with the cursor to isolate the precise structure (Figure 2B). Objects may be hidden by unchecking the corresponding box. New objects can be added as many times as necessary to refine the specificity of the structure and reduce spatial background noise. Once background noise is traced, these features may be deleted by deleting the “Object” with which they are affiliated. The desired object should be the only “Object” (all others deleted) (Figure 2C) at the end of this process, at which point one must click File → Save Object Map → Current Directory.
9. Click BMA on the right-hand side of the main panel. Select the Excel file in the folder to open the pre-generated phantom plot. Input the SigmaCT (slope) and BetaCT (offset) into the BMD scaling parameters (Figure 2D).
10. Choose Initial Segmentation, click Initial Object Map, and find the saved file of the desired object. Select this file under the Bone Object drop-down menu. Note: This should be the only object available for input since it is the only one saved. Click on Segment Cortex and Segment Trabeculae. Select Measure Bone, utilizing all default settings for this action (Figure 2E).
11. With the cursor, click and drag the image planes along the longitudinal axis of the body to grossly assess the anatomical orientation of the cortex and trabeculae by ensuring the corresponding colors for each respective bone type overlap with the correct area on the image consistently. If the color borders are inaccurate, the Cortex Threshold or Trabecular Threshold may be adjusted until proper boundaries are accomplished (Figure 2E).
12. Check the All Measures tab, which shows numerical values for all structural metrics. These measurements are automatically saved in the .CSV file within the Current Directory (Figure 2E).
D. Micro-CT quantification of lean tissue (muscle) and adipose tissue
1. Upload the mouse data and start the software applications in the same manner as outlined in section B.
2. Select an image file for loading, then double-click to open and enlarge the image. All captured images should have a unique file name including a signature sample ID, sample name, date of birth, sex, and weight.
3. Open Analyze 14.0 on the Desktop and click New Workspace. Select Input on the right-hand side. Manually search for the identical sample ID number.
4. Find the corresponding file opened previously in the Quantum GX2 Database based on the file ID number. The .VOX file should be selected for file type. Then, click Calculate it.
5. Once the image appears in the designated workspace, click Load Volume in the bottom left-hand corner of the screen. Click Exit, then Rename (if desired).
6. Select Transform on the right-hand side to localize the region of interest within the image. Choose Spatial Transforms and the Subregion function within, then click Extract Sub-Volume to compute the new volume. Click Save Volume in the lower left-hand corner of the screen.
7. Select Process from the right-hand side of the screen. Click Spatial Filter for the Process Type, Median for the Filter Type, Kernel Size 3 for the Subtype, and Automatic for the Preview Type. Then, click Process Volume. Press Save Volume.
8. Select Segment. Under the Semi-Automatic tab, click Threshold Volume. Adjust the upper and lower limits of the Threshold range until only the well-circumscribed hypodensity regions of the 3D scans are within range (depicted in red).
Note: This may be verified further by literature discussing appropriate numerical HU values for adipose density.
9. Move the cursor along one of the scans to shift the positional viewpoint within the axial, coronal, and sagittal sections. Ensure that the Threshold Rendering function is checked. Then, click Threshold Object (Figure 3A).
10. Noise can be removed by choosing Add Object. Click the Manual tab, select Manual Trace, and trace along the scan to encircle background noise within the new object domains to delineate between the body of the mouse and external structures of similar density detected by the software.
a. For example: The lung has a similar HU mean as adipose tissue. To exclude the lung from adipose, from semi-automatic, select Object Extractor, and Select Seed in the lung region. Adjust the threshold so that the whole lung is automatically selected by the Object Extractor. Select Extract Object (Figure 3B) to assign the lung to the new Object.
b. To exclude the mouse holder that has a similar HU mean as adipose tissue, go to Manual, select Polygon Trace or Manual Trace, and manually select the mouse holder to a new object (Figure 3C). Once all noise is included in the new Object, delete this Object by clicking the corresponding box and trashcan icon [Figure 3C (see the inset in the white box)]. Re-title the original Object as Fat.
Figure 3. Adipose tissue segmentation and quantification. The 3D volume rendering screenshot shows that the object extractor and threshold volume tools were used to separate fat mass. A–C. Process of separating adipose: A) threshold to segment adipose tissue; B) use Object Extractor to exclude the lungs; C) use Manual Trace tool to remove the sample bed.
11. To isolate the lean tissue, click Add Object and adjust Threshold within Threshold Volume under the Semi-Automatic tab so that the filled regions within the threshold range show color complementary to the fat and exclude bone. Check and uncheck the objects for fat and lean tissue to assess that the borders of each threshold range are non-overlapping and identifiable. Move the cursor along one of the scans to shift the positional viewpoint within the axial, coronal, and sagittal sections. Ensure that the Threshold Rendering function is checked. Then, click Threshold Object (Figure 4A).
Figure 4. Lean tissue segmentation and quantification. The object extractor and threshold volume tools can be used to separate lean mass, as seen in the 3D volume rendering screenshot. A–C: Process of separating lean tissue: A) threshold to segment lean tissue; B) use Object Extractor to exclude brain; C) use Manual Trace to remove the sample bed; the image in the inset shows the merged 3D rendering of lean and adipose tissue. D) Coronal view of the mouse micro-CT image, segmented adipose tissue, segmented lean tissue, and all three images merged. E) Screenshot of measure tool to quantify the adipose and lean mass volume.
12. Noise can be removed by choosing Add Object. Click the Manual tab, select Manual Trace, and trace along the scan to encircle background noise within the new object domains to delineate between the body of the mouse and surrounding pieces of equipment, as well as external structures of similar density detected by the software.
a. For example: The brain has a similar HU mean as lean tissue. To exclude the brain from lean tissue, from semi-automatic, select Object Extractor and select seed in the brain region. Adjust the Threshold so that the Object Extractor automatically selects the whole brain. Select Extract Object (Figure 4B) so the brain can be assigned to the new Object.
b. To exclude the mouse holder (which has a HU mean similar to lean tissue), from Manual, select Polygon Trace or Manual Trace, and manually assign the mouse holder to a new object (Figure 4C). Once all noise is included in the new Object, delete this Object by clicking the corresponding box and trashcan icon [Figure 4C (see the inset in the white box)]. Re-title the original Object as lean tissue. The merged view of segmented lean and adipose tissue is shown in Figure 4D.
13. Click File → Save Object Map to save the object map to the current directory. Click the X in the upper right corner to exit.
14. Click Measure on the right-hand side.
15. Click File → Load Object Map → Current Directory → select the file just saved and click to open.
16. Under 3-D tab, click Enabled Objects; under Stats to View, click the size intensity tab, and select Volume without changing other default settings.
17. Click Auto Log Stats and select Sample Enabled Objects in the Sample Options section (Figure 4E).
18. Copy the volumes generated for both fat and lean tissue underneath the scanned image and paste them into an Excel sheet for further calculation to obtain the percentage of body fat and lean tissue (see Data analysis).
E. Micro-CT scanning of the lungs with respiratory gating
1. Anesthetize the mouse with 2.5% isoflurane in the RAS-4 rodent anesthesia system induction chamber.
Note: It takes 2 min to fully anesthetize the mice.
Critical: For successful respiratory gating, the time between breaths should be >1,000 ms. Thus, observe the mouse during the anesthesia and ensure that the breathing is stable and not too fast.
2. Place anesthetized mice on the mouse bed and close the imaging system door. Adjust the scan settings in the control window as follows: 90 kV; current: 88 µA; acquisition: 36; recon: 36; scan mode: gating 4 min; display settings: off; X-ray filter: Cu 0.06 + Al 0.5.
Note: No stitching is necessary in this section because one scan at FOV 36 mm is sufficient to capture the chest area of an adult mouse (Figure 5A).
Figure 5. Micro-CT acquisition of mouse lungs. A, B. Video image pop-up screen of the mouse in the Quantum GX2 scanner, with the blue square indicating the FOV for the scan and the green rectangle (dotted) placed partially over the diaphragm for respiratory gating. A) Respiratory interval of less than 1,000 ms, which is incorrect. B) Corrected respiratory interval of more than 1,000 ms. C) Image shows the axial view of the mouse lungs.
3. On the control panel, select the high speed scan mode and check the respiratory gating icon in the gating technique panel.
4. Start Live Mode and move and resize the green ROI rectangle so that the moving diaphragm is covered by the ROI rectangle. Wait until the breathing is stable and the time between breaths is longer than 1,000 ms (Figure 5B, C).
5. Start CT scan. After the scanning, click Reconstruct to reconstruct the images of the inspiratory and expiratory phases.
F. Micro-CT quantification of lung volume and fibrotic structures
1. In the Quantum GX2 database application, select the image file to be loaded, then double-click to open and enlarge the image. All captured images should have a signature sample ID, sample name, date of birth, gender, and weight.
2. Launch the Analyze 14.0 application on the desktop and click Create New Workspace. Select the input function on the right. Users must manually search for the same sample ID number.
3. Find the corresponding file opened before in the Quantum GX2 Database. Then click the .VOX file to ascertain file type and select the Calculate It button.
4. After the image appears in the specified workspace, click Load Volume in the lower-left corner of the screen. Click Exit, then click Rename. Then, specify the file that needs analysis according to the preferred naming system.
5. To crop the image, use the Transform tab and click Subregion, then click on Extract Subvolume to remove the excess parts and retain only the section with the lung image. Afterward, click Save Volume, rename the file, and add “_transform” to it.
6. Select Segment and click the Semi-Automatic tab. Select Object Extractor and select seed in the lung region. Adjust the Threshold so that the Object Extractor automatically selects the whole lung. Simultaneously, one can manually input threshold values to adjust the selection range here, using HU values from -1294.1 to 0 (13) as the criteria for delineating the lung area (depicted in red) (Figure 6A).
Figure 6. Highlighting and thresholding of lung volume and fibrotic structures using Analyze software. The red areas represent non-aerated regions, indicative of fibrotic tissue. The blue areas represent normally aerated regions, reflecting healthy, well-ventilated lungs. The yellow areas correspond to hypo-aerated regions, reflecting moderate fibrosis. A) This screenshot shows how to draw ROI around the fibrosis area using the object extractor tool and adjust the binary threshold values to display the lung with a red outline. B, C) Use the Threshold Volume tool to segment normal breath and high-density areas. The blue region is the high-aerated lung area, and the red region is the high-density area in the lungs. D) Image shows that the Measure tool was used to calculate the volume of each object.
7. Lock the original image, select the semi-automatic button, use the Threshold Volume key to set the threshold from -1294.1 to -435, and frame the normal breathing image (depicted in blue) (Figure 6B).
8. Continue to select the semi-automatic button, use the Threshold Volume key to set the Threshold from -121 to 0, and frame the high-density area (depicted in red) (Figure 6C).
9. Click File → Save Object Map to save the object map to the current directory. Click the X in the upper right corner to exit.
10. Click Measure on the right-hand side.
11. Click File → Load Object Map → Current Directory → select the file just saved and click to open.
12. Under the 3-D tab, click Enabled Objects. Under the Stats to View, click Size Intensity, and select options such as Volume, Histogram, Mean Sum Entropy Number of Voxels, Std. Dev, etc.
13. Click Auto Log Stats and select Sample Enabled Objects in the sample options section (Figure 6D).
14. Segmentation validation: Compare the segmented areas (non-aerated, aerated, and moderately aerated regions) with the original micro-CT images (Figure 7A–F).
Pause point: Focus on the regions highlighted by the red circles to confirm that the segmented areas correctly represent the corresponding severity of lung fibrosis.
Figure 7. Micro-CT images of normal and fibrotic mice lungs. A–C) Axial, coronal, and sagittal lung CT images of the wild-type control C57BL/6 mouse. D–F) Axial, coronal, and sagittal lung CT images after bleomycin-induced lung fibrosis at day 21. The red circle shows the high-density area of the bleomycin-induced fibrosis mouse lungs and the normal lung area in the control mouse lungs.
Data analysis
Copy the volume generated for both “fat” and “lean tissue” underneath the scanned image and paste this into an Excel sheet (convert from mm3 to cm3). Manually input the known densities for each (0.95 g/cm3 for soft tissue and 1.06 g/cm3 for adipose and lean tissue) [11]. Multiply the volume by density to obtain the mass (g). Then, divide the mass obtained for these values by the body weight to obtain the percentage of body fat and lean tissue.
Data analysis was performed in GraphPad Prism to calculate statistical significance (n = 5–7 biological replicates) using the Student’s unpaired t-test [12]. A p-value of <0.05 was considered statistically significant. Familiarity with GraphPad Prism is necessary.
Validation of protocol
BMD calculation validation: The BMD values obtained from the calculation of the micro-CT scan of the femur cortical bone of WT mice were between 749.4 and 828.47 HA/cc. These values align with the range previously reported by Mohan et al. (845.4 ± 60.2 HA/cc) [13]. The consistency with previously reported data supports the accuracy of our BMD measurements.
Fat and lean tissue calculation validation: To validate the accuracy of fat and lean tissue quantification, we scanned a WT male mouse using micro-CT and dissected out the abdominal fat and muscle from the hind limbs. The dissected fat and lean tissue were weighed, and the measured weights (fat: 1.169 g, lean: 1.385 g) matched the values calculated from the micro-CT scan (fat: 1.349 g, lean: 1.547 g). The challenges of perfect dissection can explain the differences. This result supports the accuracy of the micro-CT quantification for fat and lean tissue.
Fibrosis scan validation: To validate the accuracy of fibrosis quantification, we performed a micro-CT scan and H&E staining on the same lung tissue samples. The micro-CT results showed that mice with lower aerated lung volumes had higher levels of fibrosis, as evidenced by the increased fibrotic areas in the corresponding H&E staining. This correlation between micro-CT and histological analysis supports the accuracy of lung fibrosis measurement using micro-CT.
General notes and troubleshooting
General notes
1. Experimental design. The Quantum GX2 micro-computed tomography scanner, which is commercially available, is used in this protocol to optimize image acquisition parameters for low radiation dose, high-resolution, and high-throughput computed tomography imaging. This methods paper presents a simplified workflow for acquiring and analyzing micro-CT images for mouse body composition, BMD, and mouse lungs by comparing normal and fibrotic lungs. It takes 4–8 min to acquire an image of each animal and 10–30 min to analyze the data. Researchers should be able to use this approach in various mouse models for research purposes if they have a basic understanding of medical imaging, animal care, and software analysis.
2. Imaging workflow. The Quantum GX2 is an advanced micro-CT imaging system (Figure 1A) designed for pre-clinical research across a wide range of applications. The RAS-4 Rodent anesthesia system (Figure 1B) connected to the Quantum GX2, where the animal can be kept anesthetized for several minutes (Figure 1C), is ideal for non-invasive, longitudinal studies, producing high-quality images (Figure 1D). The instrument has four fields of view ranging from 18 to 86 mm, with the highest resolutions of 2.3 µm voxels. Quantum GX2 is ideal for imaging pre-clinical animals, including mice, rats, rabbits, zebrafish, and ex vivo samples. This system is also capable of two-phase retrospective respiratory and cardiac gating.
3. Radiation dosing. The X-ray radiation dose is an important consideration in micro-CT application. Although micro-CT provides high-resolution images in a non-invasive manner that can be valuable in longitudinal studies, X-ray exposure can potentially affect both DNA repair and the cell cycle in mice [14]. Depending on the strain, the LD50/30 for mice is approximately 5–7.6 Gy [15]. However, the radiation dosage for a respiratory-gated lung scan is about 926 mGy. Thus, to avoid damage caused by radiation, the mouse should not be scanned too frequently. For reference, the mice utilized in this study were safe at a rate of two scans per month. Parameters in micro-CT scanning influencing X-ray dose include the exposure time, type of filters, voltage, and current. In longitudinal studies, researchers should optimize these parameters to minimize the radiation dose while obtaining high-quality images [14,15].
Troubleshooting
Problem 1: The prescribed stepwise quantification does not account for differences in adipose tissue.
Possible Cause: Similar density values between subcutaneous and visceral fat cause a lack of distinction between these tissue subtypes after micro-CT scanning acquisition.
Solution: The only method by which such tissue types may be separated would be through manual segmentation based on known anatomical location. Figure 1 in the work by Judex et al. demonstrates manual segmentation for this purpose [14], which is not discussed in this article.
Other limitations:
1. The high cost of micro-CT can limit accessibility, especially for smaller research labs or institutions with limited funding.
2. The learning curve associated with micro-CT technology may lead to inconsistencies in data quality and interpretation, particularly among less experienced users.
3. Without adequate training resources, researchers may struggle to fully utilize micro-CT technology's capabilities or produce reliable, reproducible results. This could lead to discrepancies in data quality.
4. Animal handling and animal positioning could impact the image quality and measurements.
Supplementary information
The following supporting information can be downloaded here:
1. Figure S1. Gain and HU calibration
2. Figure S2. Nose cone setup for isoflurane anesthesia delivery using the RAS-4 system
3. Figure S3. This screenshot image shows the settings for creating a new Job scan with five stitches
4. Figure S4. BMD calibration using phantom
Acknowledgments
We thank representatives and manufacturers from PerkinElmer who responded to our requests for information on their micro-CT GX2 scanner and previous works as outlined in the Validation of protocol from which our methods were derived. This research work was supported by the Lavin-Bernick grant and Tulane Center of Excellence: Sex-based Biology and Medicine (TCESBM) pilot award to M.A.
Competing interests
All authors of the present manuscript have no competing interests concerning the research conducted and presented in this manuscript, including but not limited to paid employment or consultancy, stock ownership, patent applications, personal relationships with individuals involved in the submission or evaluation of a protocol, and receipt of funding or free products from the vendors of the reagents/equipment or other advertisers.
Ethical considerations
The animals used for micro-CT analysis have been approved by the Institutional Animal Care and Use Committee Office (IACUC), Tulane University School of Medicine.
References
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Nasrazadani, A., Thomas, R. A., Oesterreich, S. and Lee, A. V. (2018). Precision Medicine in Hormone Receptor-Positive Breast Cancer. Front Oncol. 8: e00144.
Fang, K., Li, Q., Wei, Y., Zhou, C., Guo, W., Shen, J., Wu, R., Ying, W., Yu, L., Zi, J., et al. (2021). Prediction and Validation of Mouse Meiosis-Essential Genes Based on Spermatogenesis Proteome Dynamics. Mol Cell Proteomics. 20: 100014.
da Silva-Buttkus, P., Spielmann, N., Klein-Rodewald, T., Schütt, C., Aguilar-Pimentel, A., Amarie, O. V., Becker, L., Calzada-Wack, J., Garrett, L., Gerlini, R., et al. (2023). Knockout mouse models as a resource for the study of rare diseases. Mamm Genome. 34(2): 244–261.
Zaw Thin, M., Moore, C., Snoeks, T., Kalber, T., Downward, J. and Behrens, A. (2022). Micro-CT acquisition and image processing to track and characterize pulmonary nodules in mice. Nat Protoc. 18(3): 990–1015.
Ketten, D. R., Simmons, J. A., Riquimaroux, H. and Simmons, A. M. (2021). Functional Analyses of Peripheral Auditory System Adaptations for Echolocation in Air vs. Water. Front Ecol Evol. 9: e661216.
Clark, D. and Badea, C. (2021). Advances in micro-CT imaging of small animals. Physica Med. 88: 175–192.
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Beaucage, K. L., Pollmann, S. I., Sims, S. M., Dixon, S. J. and Holdsworth, D. W. (2016). Quantitative in vivo micro-computed tomography for assessment of age-dependent changes in murine whole-body composition. Bone Rep. 5: 70–80.
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Article Information
Publication history
Received: Sep 17, 2024
Accepted: Dec 16, 2024
Available online: Jan 19, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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5,208 | https://bio-protocol.org/en/bpdetail?id=5208&type=0 | # Bio-Protocol Content
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This is an In Press version of the protocol that has not yet been assigned to an issue.
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Streamlined Quantification of p-γ-H2AX Foci for DNA Damage Analysis in Melanoma and Melanocyte Co-cultures Exposed to FLASH Irradiation Using Automated Image Cytometry
SO Stefana Orobeti *
ID Ioana Dinca
AB Alexandra Bran
IT Ion Tiseanu
FS Felix Sima
SP Stefana M. Petrescu
LS Livia E. Sima *
(*contributed equally to this work)
In Press, Available online: Jan 19, 2025
DOI: 10.21769/BioProtoc.5208 Views: 40
Reviewed by: Ivonne Sehring Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Scientific Reports Jun 2024
Abstract
In response to DNA-damaging physical or chemical agents, the DNA damage repair (DDR) pathway is activated in eukaryotic cells. In the radiobiology field, it is important to assess the DNA damage effect of a certain irradiation regime on cancer cells and compare it to the effect on non-transformed cells exposed to identical conditions. The first step in the DNA repair mechanism consists of the attachment of proteins such as the phosphorylated histone γ-H2AX (p-γ-H2AX) to DNA double-strand breaks (DSB) in the nucleus, which leads to the formation of repairing foci. Therefore, imaging methods were established to evaluate the presence of foci inside the nucleus after exposure to DNA-damaging agents. This approach is superior in sensitivity to other methods, such as the comet assay or the pulsed-field gel electrophoresis (PFGE), that allow direct detection of cleaved DNA fragments. These electrophoresis-based methods require high ionizing radiation dosages and are difficult to reproduce compared to imaging-based assays. Conventionally, the number of foci is determined visually, with limited accuracy and throughput. Here, by exploring the effect of laser-plasma accelerated electrons FLASH irradiation on cancer cells, we describe an image cytometry protocol for the quantification of foci with increased throughput, upon large areas, with increased precision and sample-to-sample consistency. It consists of the automatic scanning of fluorescently labeled cells and using a gating strategy similar to flow cytometry to discriminate cells in co-culture based on nuclei elongation properties, followed by automatic quantification of foci number and statistical analysis. The protocol can be used to monitor the kinetics of DNA repair by quantification of p-γ-H2AX at different time points post-exposure or by quantification of other DNA repair proteins that form foci at the DNA DSB sites. Also, the protocol can be used for quantifying the response to chemical agents targeting DNA. This protocol can be performed on any type of cancer cells, and our gating strategy to discriminate cells in co-culture can also be used in other research applications.
Key features
• Analysis of DNA-damage sensitivity using model cancer cell lines and non-transformed cellular controls.
• Allows comparative testing of various doses of DNA damaging radiation on cancer and non-transformed cells in co-culture, as well as in monocultures.
• This protocol requires TissueFAXSiPlus model i12 or an alternative instrument that allows automatic image acquisition and stitching to benefit from enhanced analysis throughput.
• For analyses of co-cultures or heterogeneous samples, TissueQuest software is required to selectively quantify different cell subpopulations; dedicated training is advisable before operating the system.
Keywords: Cancer cells FLASH LPA electrons cell irradiation DNA damage Immunofluorescence γ-H2AX foci Image cytometry
Graphical overview
Workflow of cell co-culture irradiation and foci quantification analysis
Background
Very high energy electrons (VHEE), defined as electrons with energies between 150 and 250 MeV, were proposed as an alternative radiotherapy approach and found adequate to penetrate deep tumor sites [1]. High-intensity laser-driven acceleration of VHEE of dose rates as high as 1013 Gy/s is achievable by laser-plasma accelerators (LPA) with quasi-monoenergetic beams, being proposed as new platforms for VHEE-induced FLASH irradiation [2,3]. We have recently confirmed the cell DNA damage–producing capacity of the laser-driven high-energy dose electrons produced by a tabletop high-power LPA in cancer cells for future FLASH-radiotherapy applications [4].
In order to investigate the potential sparing effect of FLASH radiation on normal cells while evaluating the conditions where it induces enhanced DNA damage to cancer cells, we set up a protocol that allows simultaneous exposure of normal and malignant cells to irradiation by LPA-produced electrons for comparison. The golden standard method in the field of radiobiology to evaluate the effects of radiation on cells is by analyzing nuclear foci that accumulate at DNA damage sites [5,6]. The ionizing radiation is known to produce DNA double-strand breaks (DSBs) that affect preponderantly the cells engaged in the cell cycle [7]. Therefore, this type of radiation is used in radiotherapy to eliminate highly proliferative cancer cells from tumors [8]. Upon exposure of cells to radiation or other DNA-damaging agents, the DNA repair molecular mechanism is activated [7]. Proteins accumulating and forming foci at the sites of insult can be used as markers to quantitate the extent of the produced damage [6], as well as for monitoring the kinetics of DNA repair [9]. Ideally, the radiation regimen should have a maximal impact on cancer cells without affecting the normal function of other cells in the tissue. Using our proposed co-culture protocol, one can simultaneously observe the effects produced by a certain radiation dose and regimen on other cells in the target tissue alongside the cancer cells. This approach has several key advantages, namely 1) minimizing experiment-to-experiment variability when comparing the effect of a certain irradiation regime (type of source and dose) on different cell types; 2) allowing cell–cell communication between different cell types that modulate the outcome on each cell type, similarly to events produced within the tumor microenvironment; and 3) minimizing the time required for radiation exposure, thereby increasing feasibility of large experiments where several doses have to be applied in replicates in the same day on cells from the same batch. Our method allows discriminating between different cell types within the same sample based on morphological and fluorescence parameters (image cytometry). Here, we present a case where different cell types have different nuclear shapes [A375 cells have round nuclei, while normal human epidermal melanocytes (NHEMs) have elongated nuclei]. Nevertheless, TissueQuest enables users to separate cell populations based on protein markers’ expression or other features that can be quantified using fluorescent labeling.
For this, we have harnessed the power of quantitative imaging that allows precise evaluation of foci number with increased throughput on a statistically significant number of cells per sample. Using an automatic scanning system and image cytometry software, we have recently compared the effects of VHEE radiation and conventional X-ray in co-cultures of melanoma (A375 cells) and NHEMs [4]. Upon recovery and labeling of irradiated cells with antibodies against p-γ-H2AX and a nuclear stain (i.e., Hoechst), we have successfully shown that the accelerated VHEE can induce an increase in DNA damage foci count in cancer cells. Noteworthy, in a specific area of the co-culture monolayer, the effect is only observed in cancer cells, while spearing the NHEMs from stress [4].
The protocol presented here can be adopted by researchers studying the effect of any type of radiation on adherent cells, as well as by investigators who apply other types of DNA damage agents to cells in vitro (e.g., cisplatin). Also, the detailed presentation of the quantitative cytometry analysis strategy will be useful for TissueFAXS microscopy users to implement automatic scanning and analysis in their laboratories.
Materials and reagents
Biological materials
1. A375 human metastatic melanoma cells [American Type Culture Collection (ATCC), catalog number: CRL-1619)]. The cells should be grown to 70%–80% confluency in A375 culture media (see Recipe 1) and kept in a humidified environment at 37 °C, in the presence of 5% CO2, in a cell culture incubator
2. Normal human epidermal melanocytes (NHEMs) (Lonza, catalog number: CC-2504)
Reagents
1. High-glucose (HG) Dulbecco’s modified Eagle’s medium (DMEM) containing L-GlutaMAX (Gibco, catalog number: 61965-026)
2. Melanocyte growth basal medium 4 (MBM-4) (Lonza, catalog number: CC-3250)
3. Melanocyte growth medium 4 (MGM-4), Single Quots Supplements (Lonza, catalog number: CC-4435)
4. Fetal bovine serum (FBS) (Gibco, catalog number: 10437-028)
5. Penicillin-Streptomycin, 10,000 U/mL to 10,000 μg/mL (Gibco, catalog number: 15140122)
6. 1× Dulbecco's phosphate-buffered saline (DPBS) without calcium (Ca) and magnesium (Mg) (Gibco, catalog number: 14190250)
7. 0.05% Trypsin-EDTA solution (Gibco, catalog number: 25300054)
8. 0.025% Trypsin-EDTA solution (Lonza, catalog number: CC-5012)
9. 0.02% Versene (EDTA) solution (Lonza, catalog number: 17-711E)
10. Trypsin neutralizing solution (TNS) (Lonza, catalog number: CC-5002)
11. 0.4% Trypan Blue solution (StemCell Technologies, catalog number: 7050)
12. Bovine serum albumin (BSA) powder (Sigma-Aldrich, catalog number: A9647-10G)
13. Cisplatin powder (Santa Cruz, catalog number: sc-200896)
14. Mouse monoclonal antibodies for the Ser140 phosphorylated form of γ-H2AX (Invitrogen, Thermo, catalog number: MA1-2022)
15. Alexa Fluor 594-conjugated donkey anti-mouse secondary antibodies (Invitrogen, Thermo, catalog number: A21203)
16. Triton X-100 solution (Sigma-Aldrich, catalog number: T9284-500ML)
17. Paraformaldehyde (PFA) powder (Sigma-Aldrich, catalog number: P6148-500G)
18. Hoechst 33342 (Molecular Probes, Thermo, catalog number: H21492)
19. FluorSave Reagent (Millipore, catalog number: 345789)
20. NaOH (ADRA CHIM SRL, catalog number: 011-002-00-6)
Solutions
1. A375 cell culture media (complete DMEM HG) (see Recipes)
2. NHEM cell culture media (complete MBM-4) (see Recipes)
3. 10 mM cisplatin stock solution (see Recipes)
4. 5 M NaOH solution (see Recipes)
5. 4% PFA solution (see Recipes)
6. 0.2% Triton X-100 permeabilization solution (see Recipes)
7. 0.5% BSA blocking solution (see Recipes)
8. Primary antibody incubation solution (see Recipes)
9. Secondary antibody incubation solution (see Recipes)
10. Hoechst staining solution (see Recipes)
Recipes
1. A375 cell culture media
Reagent Final concentration Quantity or Volume
DMEM HG medium n/a 445 mL
Heat-inactivated FBS 10% 50 mL (see note*)
Penicillin-streptomycin solution 100 units/mL penicillin, 100 μg/mL streptomycin 5 mL
Total (optional) n/a 500 mL
*Note: Before preparing the media, inactivate the bottle of FBS into a water bath for 30 min at 56 °C. Allow the heat-inactivated FBS (hiFBS) solution to cool down before adding it to the complete DMEM HG media. Prepare hiFBS aliquots and store them in the -25 °C freezer.
2. NHEM cell culture media
Reagent Final concentration Quantity or Volume
MBM-4 basal medium n/a 500 mL
MGM-4 Single Quots Supplements n/a 9 mL
Total (optional) n/a 509 mL
*Note: After adding MGM-4 Single Quots Supplements to MBM-4 basal medium, use within one month. Do not re-freeze.
3. 10 mM cisplatin stock solution
Reagent Final concentration Quantity or Volume
Cisplatin powder n/a 30 mg
MilliQ ultrapure water n/a Adjust to 10 mL
Total (optional) 10 mM 10 mL
4. 5 M NaOH solution
Reagent Final concentration Quantity or Volume
NaOH pellets n/a 20 g
MilliQ ultrapure water n/a Adjust to 100 mL
Total (optional) 5 M 100 mL
5. 4% PFA solution
Reagent Final concentration Quantity or Volume
PFA powder 4% 2 g
PBS (see note *) 1× Adjust to 50 mL
NaOH solution n/a 3 μL sequential pipetting/dropwise until a clear solution is formed (~pH 7)
Total (optional) n/a 50 mL
*Note: Heat 40 mL of PBS in a glass beaker under a fume hood to ~60 °C. Add the PFA powder and stir until it dissolves. Cool the solution to room temperature (RT) and then adjust the pH to 7 using NaOH. Bring the volume to 50 mL. Aliquot and store in the freezer.
6. 0.2% Triton X-100 permeabilization solution
Reagent Final concentration Quantity or Volume
Triton X-100 solution (see note *) as produced (0.2–0.9 mM) 100 μL
PBS solution 1× 50 mL
Total (optional) 0.2% 50 mL
*Note: Triton X-100 is a viscous solution. Cut the pipette tip before aspirating the solution. Release the tip in the PBS solution after resuspending it three times.
7. 0.5% BSA blocking solution
Reagent Final concentration Quantity or Volume
BSA powder n/a 0.25 g
PBS solution (see note *) 1× Adjust to 50 mL
Total (optional) 0.5% 50 mL
*Note: Add small amounts of PBS solution onto the BSA powder and mix gently. Otherwise, the mixed solution will become foamy. If so, centrifuge at the maximum speed rotation allowed for the centrifuge tube in which you dissolved the powder.
8. Primary antibody incubation solution
Reagent Final concentration Quantity or Volume
Mouse anti-p-γ-H2AX 1 mg/mL
BSA blocking solution 0.5%
Total (optional) 1:200
9. Secondary antibody incubation solution
Reagent Final concentration Quantity or Volume
AlexaFluor 594-conjugated donkey anti-mouse secondary antibody 2 mg/mL
BSA blocking solution 0.5%
Total (optional) 1:400
10. Hoechst staining solution
Reagent Final concentration Quantity or Volume
Hoechst 33342 stock solution 10 mg/mL 1 μL
PBS solution 1× 3 mL
Total (optional) 1:3,000 3 mL
Laboratory supplies
1. T-75 cell culture flasks (Corning Life Sciences, catalog number: 430641U)
2. T-25 cell culture flasks (Corning Life Sciences, catalog number: 430639)
3. SlideFlasks (Thermo Scientific, catalog number: 734-2107); the package contains a chamber detachment tool
4. Sterile Corning 24-well polystyrene tissue culture plates (Corning Life Sciences, catalog number: 3526)
5. Sterile 15 mL polypropylene centrifuge tubes (Corning Life Sciences, catalog number: 430790)
6. Sterile 50 mL polypropylene centrifuge tubes (Corning Life Sciences, catalog number: 430828)
7. 2 mL serological pipettes (Corning Life Sciences, catalog number: 4486)
8. 5 mL serological pipettes (Corning Life Sciences, catalog number: 4487)
9. 10 mL serological pipettes (Corning Life Sciences, catalog number: 4488)
10. Microcentrifuge tubes (ratiolab, catalog number: 5616027)
11. 12 mm diameter coverslips (CS), circular (Marienfeld, catalog number: 0111520)
12. Microscope slides (Marienfeld, catalog number: 1000200)
13. Forceps (Millipore, catalog number: XX6200006P)
14. Syringe needles
15. Pipette tips (compatible with your automatic pipettes)
16. Microscope cover glasses for chamber slides (Nunc/Thermo, catalog number: 171080)
17. Marienfeld counting chambers (Neubauer, catalog number: MARI0640130)
18. Microscope cover glasses 22 × 22 mm for counting chamber (Paul Marienfeld GmbH & Co, Marienfeld SUPERIOR, catalog number: 0101050)
19. Carl Zeiss Immersol immersion oil for microscopy (Zeiss, catalog number: 518F)
Equipment
1. Cell culture incubator maintained at 37 °C and 5% CO2 (Thermo Scientific, model: Heracell VIOS 250i, catalog number: 13-998-253)
2. Laminar-flow biosafety cabinet (Thermo Scientific, model: KS 15, Class II)
3. Refrigerated centrifuge (Beckman Coulter, model: Allegra X-12R)
4. MilliQ water purification system (Millipore, Model SAS 67120 water purification with filter Biopak Polisher, catalog number: CDUFBI0A1)
5. Stripettor Ultra Pipet Controller (Corning, catalog number: 15536304)
6. Eppendorf Research plus P10 mechanical pipette (Eppendorf, catalog number: 3123000020)
7. Eppendorf Research plus P20 mechanical pipette (Eppendorf, catalog number: 3123000039)
8. Eppendorf Research plus P200 mechanical pipette (Eppendorf, catalog number: 3124000083
9. Eppendorf Research plus P1000 mechanical pipette (Eppendorf, catalog number: 3123000063)
10. X-ray system consisting of a NIKON XT H 225kV reflection target source with 3 μm focal spot size (Nikon Metrology NV) and a PTW UNIDOS T 10005-50406 Electromer connected to a Farmer ionization chamber TN30010 (calibrated at PTB Germany)
11. X-ray detector system: Gafchromic EBT3-810 dosimetry film (Ashland Global Holdings Inc., product code: 828204)
12. Customized chopper manufactured by selective laser melting (SLM) using 3D printing from steel with brass infusion. X-ray chopper specifications: 12 slits of 8 mm (height) × 1.5 mm (width) at 15° apart from each other
13. LPA system (Thales, France) based on a Ti:Sapphire high-intensity laser (800 nm, 25 fs pulse duration, 25 J energy at 0.1 Hz, 1 J at 10 Hz), beam transport (Ardop, France), interaction chamber (Astra, Gemini, RAL UK)
14. Electron beam detector systems: ionization chamber (model Advance Markus, type TN34045), three thermoluminescent dosimeters (model Panasonic), Gafchromic film (model GF-EBT3)
15. Automated imaging system (TissueGnostics, Vienna, Austria, model: TissueFAXSiPlus i12), based on the Zeiss Axio Observer.Z1 motorized inverted microscope appended with an ultra-precise motorized stage for automated sample acquisition with:
a. Oil immersion objective lens (Zeiss, model: Plan-Apochromat 63×/1.4 Oil DIC M27, 0.19 mm working distance)
b. Air objective lens (Zeiss, model: LD Plan-Neofluar 10×/0.3 M27, 5.3 mm working distance)
c. Air objective lens (Zeiss, model: LD Plan-Neofluar 20×/0.4 Korr M27, 7.9 mm working distance)
d. DAPI filter (excitation λ: 360 nm; emission λ: 462 nm), associated to the DAPI channel
e. Alexa 568/Cy3 filter (excitation λ: 568 nm; emission λ: 603 nm), associated to the TxRed channel
f. High-sensitivity digital monochrome camera for fluorescence microscopy (PCO AG, Kelheim, Germany; model: PCO PixelFly 14-bit dynamic range grayscale CCD camera; catalog number: C11440-10C)
g. Microscope fluorescence light source: Excelitas X-Cite 120PC Q (Cambridge Scientific, catalog number: 11419)
Note: Full instrument configuration is available at https://www.biochim.ro/facility-11/.
Note: Due to the necessity of recovering the cells in a CO2 incubator before fixation (and also keeping them in a physiological milieu immediately prior to the electrons exposure), the irradiation facility should be close to a tissue culture lab. Alternatively, for short periods of time (up to 4 h), cells could be kept at 37 °C in HEPES buffered media. This should be tested before the experiment because not all cell lines cope well in these conditions.
Software and datasets
1. TissueFAXS Slides Module (version 3.5.5.0129, release date 20 August 2012)
2. TissueQuest (version 4.0.1.0140, release date 18 November 2014)
3. Microsoft Office 365 Excel
4. GraphPad Prism (version 9.5.1 (528), release date 24 January 2023)
Procedure
A. Setup of cell culture conditions for irradiation
1. Culture vessels priming before cell seeding for the irradiation experiment: One day before the experiment, incubate each SlideFlask that will be irradiated or kept as non-irradiated control with a mix of 1 mL of A375 cell culture media and 1 mL of NHEM cell culture media. Keep them for 30 min in a humidified environment at 37 °C with 5% CO2 in the cell culture incubator before plating the cells.
Note: In parallel, it is advisable to seed similar samples in wells of a 24-well plate to be treated with drugs (e.g., cisplatin at IC50 or other DNA damage-inducing concentration) that induce the desired effect, as chemical positive controls. Make sure the surface quality of the wells is similar to the one of the irradiated vessels so they can be compared: borosilicate cover glass, tissue culture plastic, glass bottom chambered slide, etc. Cells have different attachment behaviors on different surface qualities with consequences on proliferation, confluency generation, and overall functionality.
Critical: A coverslip dedicated to the secondary antibody-only negative control should be prepared for checking antibody specificity and setting the background threshold.
Note: Wait until the cell monolayers from your cell culture flasks are at approximately 80% confluency. Then, proceed with detaching and plating them in SlideFlasks for the irradiation experiment.
Note: Prior to the experimental procedure, the number of cells to be seeded should be determined in order to obtain 80%–90% confluency the next day. Monolayers should not be overconfluent at the final endpoint, because they could easily detach from the surface during the washing procedures or even die and float before fixation.
2. Cell seeding for the irradiation experiment: Culture the cells in SlideFlasks 24 h prior to the irradiation procedure. After growing your cells in specific culture vessels (e.g., T-75 cells culture flasks for A375 and T-25 flasks for NHEM cells) in the cell incubator, bring them to the laminar flow hood and proceed with cell detachment.
a. Aspirate the cell culture media from the T-75 cell culture flasks in which A375 were cultivated. Wash the cells using 10 mL of PBS. Detach the cells with 1 mL of 0.05% Trypsin-EDTA solution for 5 min in the cell incubator. Stop the trypsinization by adding 4 mL of culture media. Pipette up and down three times and transfer the 6 mL of cell suspension to sterile 15 mL centrifuge tubes. Centrifuge the cells at 200–300× g (1,200–1,500 rpm) for 5 min at room temperature (RT). Remove the supernatant and add 2 mL of culture media onto the cell pellet. Resuspend three times and harvest 500 μL of cell suspension to dilute it with 0.4% Trypan Blue solution for counting with Marienfeld cell counting chambers. Use a density of 840,000 A375 cells in 1 mL of A375 cell culture media to seed each SlideFlask for co-culture with NHEM cells.
Note: Dilute the cell suspension with the appropriate dilution factor of 0.4% Trypan Blue solution (i.e., 1:2, 1:5, 1:10) to count between 20 and 50 cells per each counting chamber 4 × 4 square field.
b. Aspirate the cell culture media from the T-25 cell culture flasks in which NHEM cells were grown. Rinse the cells gently with 5 mL of PBS at RT by pipetting it down the cell-free surface of the culture vessel. Aspirate the PBS from the flask. Add 1 mL of 0.025% Trypsin-EDTA solution diluted with 1 mL of 0.02% Versene and incubate the cells for 1 min in the cell incubator. Neutralize with 2 mL of TNS at RT and wash the flask with 2 mL of culture media. Collect all the cells and transfer them to sterile 15 mL centrifuge tubes. Centrifuge the cells at 100× g for 3 min at RT. Throw the suspension and resuspend the cell pellet with 1 mL of culture media. Dilute with 0.4% Trypan Blue solution to count the cells. Use 420,000 cells in 1 mL of NHEM cell culture media for each SlideFlask co-cultured with A375 cells.
Caution: NHEM cells disassociate easily from the flask surface; work gently with them.
B. Irradiation using LPA electrons or X-rays
1. On the day of irradiation, fill up the SlideFlasks with media and seal them by completely screwing the caps. This is necessary to prevent cell drying when the flask is tilted and accommodated in the irradiation chamber.
Note: If irradiating co-cultures, use either a mix of equal volumes of complete media of the component cell lines or a standard complete media of a cell line that does not negatively affect the other ones present in the flask.
Caution: If using vented caps, make sure the filter stays dry throughout the procedure so as not to encourage microbial infection, especially if the endpoint analysis timepoint is later than 16–18 h. Also, to prevent potential spilling of media through the vent, add parafilm around the cap.
2. For X-ray irradiation, mount the SlideFlasks vertically with double adhesive tape onto the vertical support placed in front of the beam-generating system (Figure 1A), making sure not to provoke vibrations or other mechanical stress that would detach the cells from the flask.
Pulsed X-ray setup: The setup used for delivering pulsed X-rays consists of an X-ray source and a customized X-ray beam chopper. It includes a sample holder that positions the sample in front of the head of the X-ray source. Operate the X-ray source at a voltage of 200 kV and a current of 100 μA using copper foil with a thickness of 0.2 mm as pre-filtration.
Chopper operation: Rotate the chopper at high speed using a brushless DC motor powered by an adjustable power source (24 V, 60 A) set at 19 V. Control the motor with an electronic speed controller unit. Regulate the rotational speed using an electronic board that generates a PWM (pulse width modulated) signal controlled by LabView using LINX library. Set the X-ray pulse frequency to 1.4 kHz.
Irradiation conditions: Position the sample at a distance of 10 cm from the X-ray source and irradiate with pulsed X-rays for 15 min and 30 s. To ensure optimal shielding of the X-ray beam passing through the chopper slits and to reduce the X-ray scattering, place Pb collimators in front of the X-ray source (aperture of ~4 mm, total thickness of ~0.8 cm). Carefully align the chopper slits with the collimator aperture and the center of the sample.
Note: For these operating parameters, under continuous exposure, a dose rate of 26 mGy/s at a distance of 10 cm was measured by an ionization chamber connected to a standard UNIDOS dosimeter.
Dose measurements: Use Gafchromic EBT3 films to estimate the doses delivered to the samples. Place them on the back of the SlideFlask, in front of the plastic-attached cells, as viewed from the source. Prior to experiments on cell cultures, irradiate EBT3 films with precisely delivered doses, and generate calibration curves from the radiochromic film darkening, according to Campajola [14].
Note: In our experiments, the samples irradiated with pulsed X-rays for 15 min and 31 s received a total dose of 2.4 Gy, according to Gafchromic EBT3 measurements.
Figure 1. Photographic image exemplifying the mounting of media-filled SlideFlasks onto the support before targeting the sample with the X-ray beam (A) or LPA electrons (B)
3. LPA setup: The setup used for delivering very-high-energy electrons (VHEE) consists of a high-intensity laser beam focused with an off-axis parabolic mirror with a 3.2 m focal length into a supersonic gas jet consisting of 99% He + 1% N2. Use high-intensity petawatt laser-driven accelerators that can deliver very-high-energy electrons (VHEEs) at dose rates as high as 1013 Gy/s in very short pulses (10-13 s).
Note: For monitoring the effective applied dose, you can apply various radiation dose detectors and sensors on the flask surface close to the cell monolayer (e.g., in situ dose monitoring by ionization chamber, thermoluminescent dosimeters (TLDs) for evaluating cumulative dose post-experiment, or Gafchromic films for dose uniformity check).
The quasi-monoenergetic electron energy distributions can be measured with a spectrometer equipped with a Pb collimator and magnetic dipole of 0.8 T, which can be set up into the electron beam at the extension of the interaction chamber.
As an example, for simultaneous irradiation of two containers (Figure 1B) with cells grown either in the first or both containers, introduce three TLDs: one placed in front of the first container, the second placed between the two containers, and the third placed after the second container. In our setup, the electron beam travels approximately 198 cm in vacuum before reaching the first TLD, and then it travels successively through a 60 μm Al foil, a 1 cm thick glass window, another 60 μm Al foil, a 0.5 mm cardboard, the scintillator screen LANEX, and 2 cm in air.
Note: In our experiments, the samples irradiated with VHEE received a total dose of approximately 150 mGy, according to ionization chamber dosimetry measurements.
General note: Irradiate cancer cells and normal melanocyte co-cultures with either VHEE produced by LPA or pulsed X-ray beams (for control experiments). X-ray equivalent doses of up to 8 Gy are commonly used to study radiation response on in vitro cultured cell line models [15].
C. Cell recovery and fixation
1. Upon irradiation, allow cells to recover and accumulate phosphorylated γ-H2AX [10] at the double-strand DNA breaks for 30 min in the incubator.
Note: If flasks do not have vented caps, slowly release the cap to allow gas passage into the vessel. Remove a fraction of the media to prevent overflow; leave enough media to cover the cells and consult the working volume recommended by the vessel producer.
2. Aspirate growth medium and gently wash cells once with a generous volume of PBS (at least equal to the media volume used for culture) at RT.
3. Fix cells using 4% PFA for 30 min at RT. Use a sufficient amount to cover the cell monolayer.
Caution: PFA is neurotoxic. Use a chemical hood or work in a well-ventilated space when performing the procedure. Also, all derived chemical hazardous waste should be collected separately and sent to appropriate facilities for neutralization. Do not discard solutions into the sink!
4. Aspirate fixing solution and gently wash cells twice with PBS at RT. Use at least double the amount of PFA used.
Pause point: Fixed cells can be kept in PBS at 4 °C for 1–2 weeks before immunofluorescent labeling. It is, however, advisable to proceed with the next steps immediately or within days as longer storage may reverse fixation affecting cell morphology and epitope integrity. Refill wells/flasks with PBS periodically and/or seal with parafilm to prevent cell drying.
D. Immunofluorescence labeling
1. Aspirate PBS from fixed co-culture cells plated on SlideFlasks or coverslip (CS) controls. Add a volume of 3 mL/SlideFlask and 350 μL/CS of 0.2% Triton X-100 permeabilization solution. Incubate for 3 min at RT.
2. Wash twice with PBS and incubate for 1 h at RT with 0.5% BSA blocking solution using the same volume as in the permeabilization step.
3. Before the immunostaining step, detach the flasks from the slides with the aid of the chamber detachment tool provided within the SlideFlasks package.
4. Add 350 μL/slide and 250 μL/CS of primary antibody incubation solution, respectively (except for the negative “secondary only” control, where you add blocking buffer). Place the slides in a horizontal position facing upward in a humidity chamber incubation box (any type of rectangular, transparent food box can be repurposed into a humid incubation chamber by placing wet napkins on its bottom to create a humid atmosphere; additionally, a plastic grid can be used to support the slides deposited onto the napkins and prevent slide wetting; one can also use a pipette tips box from which the tips’ support part is removed). Put the lid on the box and leave the primary antibodies to react for 30 min at RT.
Note: If using other antibody clones, please check the appropriate dilution factor and the incubation time recommended by the manufacturer in the product information sheet. If not mentioned, test them on your specific cell line.
5. Rinse three times with PBS for 5 min each step.
6. Label the cells for 20 min with secondary antibody incubation solution: add 350 μL/slide and 250 μL/CS, respectively. Incubate the slides in a humidity chamber incubation box covered beneath with wet napkins to create a humid atmosphere, as in step 4.
Critical: Include here the secondary-only negative control for demonstrating signal specificity and setting the background threshold.
Critical: At this step, cover the box with aluminum foil to prevent light from damaging the fluorophores conjugated to the secondary antibodies.
7. After three washes with PBS, incubate the cells at RT for 1 min with Hoechst staining solution.
8. Rinse three times with PBS and once with MilliQ water to prevent the formation of salt crystals.
9. Use 100 μL/slide and 15 μL/CS of FluorSave Reagent, respectively, to mount the samples with the cell culture surface onto the microscope slides.
Critical: If you use circular 12 mm CSs to grow cells into 24-well plates, make sure the CSs mounted on a microscope slide are placed in the center and not at the distal areas of the slide, where the holder edge will mask the visualization of the samples. Make sure to leave the slide edges free when mounting the CS(s) to allow the slide to fit into the microscope’s slide holder.
Pause point: Mounted slides can be kept in the dark (covered with aluminum foil or stored in a dedicated slide box/cardboard folder) at 4 °C for up to one month before image acquisition without significant signal loss. It is however advisable to proceed with the next steps immediately or within days, as longer storage may favor the development of air bubbles due to drying of the mounting solution layer. To prevent this, one can seal the coverslips with transparent nail polish.
E. Image acquisition: slides automatic scanning
1. Start up the imaging system (power source, halogen lamp, microscope, PC).
2. Open the software that controls the imaging instrument by double-clicking the specific icon on the computer desktop. Make sure the slides’ sample holder is mounted on the stage and its planarity is optimal. Calibrate the motorized stage (if available) with the proper slide holder installed.
Note: In the case of the TissueFAXSiPlus system, open the TissueFAXS Slides module on the workstation desktop and calibrate the stage for slide acquisition as prompted by the software (see General note 1).
3. Carefully fit the slides into the holder with the cell-covered surface toward the objective as for any inverted microscope visualization.
4. Start sample observation and image focusing using the objective with the smallest magnification available on your imaging instrument. The light path should be set to the ocular visualization mode, and the excitation lamp should be switched on. To easily find the correct focal plane, use the DAPI channel for cell nuclei visualization first. Then, check the signal on the TxRed channel to visualize phosphorylated γ-H2AX (p-γ-H2AX) that labels damaged DNA foci.
Note: Whenever you pause visualization of the samples, switch the lamp to standby mode to prevent photobleaching in the illuminated area of your sample.
5. Prepare the system for image acquisition. For this, set up a new project using the image acquisition software. Give the new file a name and browse to select the directory for saving and storing the data.
Note: For TissueFAXSiPlus users, make sure the right image acquisition template is selected from the software menu before creating a new experiment project, by selecting Tools/Options/Default Experiment Settings/Preview. Choose slide orientation (e.g., Label Up/Label Down). One can choose to scan the whole slide, the center part of the specimen, or a custom region created by the user. Create a new project for a generic sample [not TMA (tissue microarray)] and select a 5× objective for preview and a 63× oil objective for field of view (FOV) acquisition. Select DAPI and TxRed channels for sample imaging. Save the file in the Setup folder by creating a subfolder with your name/project and a sub-subfolder with the name of the experiment.
Critical: In the case of TissueFAXSiPlus system, make sure to define the desired location on the computer where the project file and associated image folders will be stored before sample acquisition. Also, do not perform any modifications to the data storage folders post-acquisition, such as moving the folders to a different location or renaming folders. The acquisition software (or the TissueFAXS Viewer visualization software) will not be able to retrieve the FOVs to generate stitched images of region overviews anymore!
6. Start live imaging through the camera by clicking the dedicated button in the visualization window on your computer screen. The light path should be set to camera visualization mode using a proper camera dedicated to fluorescence image acquisition.
Note: In the case of the TissueFAXSiPlus system, you should choose the PCO PixelFly camera attached on the right side of the microscope by selecting the right (R) side port in the TissueFAXS Slides software window dedicated to visualization mode. This action can also be performed by selecting the R side port from the light path specified on the Zeiss microscope digital display.
Note: If the system has more than one available digital camera, make sure you select the correct camera from the imaging software menu. In the case of the TissueFAXSiPlus system, select the camera manager tab in the Tools/Options menu and choose PCO PixelFly from the dropdown list.
7. (Optional) If you have an automated scanning system, you can perform sample preview to generate a sample map and define specific regions of interest (ROI) areas to be scanned. For this, use a low magnification objective (e.g., 5× objective) and the DAPI signal detection channel. Refine focus and use exposure parameters that allow nuclei visualization.
Note: For the TissueFAXSiPlus system, double-click the Preview tab in the top-left side of the software window and press Acquire after selecting the slide position specific to your sample (the holder may have 8 or 12 slide positions). Choose Current focus position when prompted by the dialog window.
At the end of the scanning step, define the specific areas (ROIs) to be scanned using a high-magnification objective (e.g., 63× oil objective). For this, double-click the obtained preview to generate a separate enlarged visualization window. Here, choose on the left side the desired tool to define regions with a specific geometry (e.g., rectangle, circle, custom freeform). Use the mouse to select the ROI on the previewed slide. A region will be automatically created under the selected slide. Rename “Slide x” (x is the number defining the slide position into the holder) to the name of your sample. Rename “Region 001” to the name of your ROI. Repeat the creation of ROIs for all areas to be investigated.
Critical: In the case of the TissueFAXSiPlus system, make sure to define slide and region names before acquiring! No modification is possible after the acquisition.
8. Set the exposure times specific for each channel (e.g., DAPI or TxRed) and objective magnification (e.g., 63× oil objective) using both negative and most brightly stained controls to ensure that fluorescent signals are within the dynamic range of the markers’ expression. Use the prepared secondary-only sample to set the background threshold. For p-γ-H2AX signal detection setup, use the cisplatin (CisPt)-treated cells sample to fit the bright intensity signal into the visible dynamic range without detectable overexposure. Save the set parameters for the chosen magnification and do not perform any modification within a sample set to allow objective comparison between experimental conditions.
Note: In the case of the TissueFAXSiPlus system, you can start by pressing the Auto button for exposure time for each channel and refine exposure time (ideally no more than 300 ms exposure time) and background lower and upper thresholds to optimize signal-to-noise ratios.
9. Start sample image acquisition. Use either an automating image scanning instrument (such as the TissueFAXSiPlus or any other slide scanning system) to sample representative regions (ROIs) of your samples or capture a sufficient number of images with a total number of cells >500 per sample (as in [11]) for quantitative analysis.
Note: In the case of the TissueFAXSiPlus system, go to Tools/Options and select the Focus tab in Scan Settings to select sample focusing mode (e.g., Autofocus/Manual/Current position). For the best fine-tuning, select Manual. For the most efficient scanning speed, select Autofocus and press “Set around this point” after manual optimization of focus using the microscope fine-tuning knob. Press Save and Exit to save and close the dialog window. Select the region to be acquired and right-click to open the associated drop-down menu; press Acquire to start scanning the selected region.
Caution: If using Autofocus, make sure the sample does not contain artifacts; otherwise, the focus will be incorrectly diverted toward them. After the scanning has finished, check the quality of the images carefully and correct the FOVs with non-optimal focus. This can be done by selecting the flag icon above the ROI image and clicking the FOVs to be corrected. Then, right-click to select the desired re-acquisition mode (e.g., “reacquire FOV”). Images will be overwritten so that the selected FOVs can be corrected.
Note: Images of all samples within the experiment should be acquired on the same day if possible, without modifying any of the excitation or imaging parameters (lamp excitation level, exposure time, background threshold levels, room illumination, etc.) to ensure consistency. Ideally, the microscope should be placed in a dark room for fluorescent imaging; otherwise, use blinders to block sunlight passing through the windows. Sunlight affects fluorophores conjugated to secondary antibodies. Also, do not spend too much time on a specific region on the sample during the initial (exposure time) setup to prevent photobleaching. Any variation in acquisition conditions will produce errors in signal quantitation and sample comparative analysis.
10. Repeat step E9 for all samples and regions to be analyzed.
Note: In the case of the TissueFAXSiPlus system, all images are automatically stored in the folder associated with the project as grayscale captures. For pseudocolor visualization, open the files using the acquisition software and export as .tiff or .jpeg files in color mode, as individual channel-specific images or as merged versions of the overlapped channels. One can export region overviews (Figure 2A) or specific FOVs (Figure 2B) using the Export button situated at the top of the generated scan.
Figure 2. Example of an automatic image scanning result obtained using the 63× objective. A single .aqproj file consists of images captured on the DAPI and TxRed fluorescence channels for all samples and regions in a project. One region overview (A, scale bar: 1 mm) and one field of view (FOV) (B, scale bar: 20 μm) are shown for exemplification. Separate DAPI and TxRed channel images and enlarged insets are presented in grayscale (B). The p-γ-H2AX signal localizes to the nucleus in discrete regions known as foci.
Pause point: Once images are saved and stored into folders, quantitative analysis can be performed anytime afterward.
F. Image processing: cytometry analysis for p-γ-H2AX foci quantification
Images of the positive control sample should be used as a reference to set the parameters’ cutoffs and the gates for foci quantification, using the forward and backward gating functions of the software. In the case of Orobeti et al. [4], A375 cells that were treated with 2 μM CisPt for 24 h were used to set up parameters. Examples of images taken for DAPI and TxRed signals are shown in Figure 3.
Figure 3. Example of images obtained from CisPt-treated cells and untreated controls using the 63× objective. Overlapped DAPI and TxRed FOV colored images, as well as separate grayscale DAPI and TxRed channel images, are presented. The merged images of both channels as well as the TxRed-captured images show that p-γ-H2AX signal is enhanced upon treatment with CisPt (A) as compared to the untreated cells (B). The small, elongated nuclei represent NHEM cells, while the circular bigger ones belong to A375 cells. Scale bar: 20 μm.
1. Open the TissueQuest fluorescence image cytometry software by double-clicking the shortcut icon on the desktop.
Note: For this Bio-protocol, we specifically exemplify how to use the TissueQuest image cytometry software associated with the TissueFAXS scanning instruments to automatically count and analyze nuclear foci (see General note 1). However, analysis can also be performed with other image analysis packages such as the freely available ImageJ using a particle analysis plugin to quantify the foci with each image being processed separately [12]. Inversely, one can acquire separate images or region overviews with other fluorescence microscopes or automatic scanners and upload them into TissueQuest for quantitative analysis.
2. In the opened dialog window (see Figure 4), select the option “Import a TissueFAXS project” and choose the .aqproj project file from the specific experiment folder. This will initiate ROI image data uploading into the analysis software.
Note: If you want to continue or modify an already initiated quantitative analysis, select the option “Open an existing TissueQuest project.” If you want to analyze samples imaged with other microscope systems, choose the appropriate selection corresponding to the instrument you used (e.g., Mirax Scanners, Panoramic Scanners, or Zeiss Scanners).
Figure 4. Screen capture of the “TissueQuest Open/Import” dialog window
3. In the Choose TissueFAXS Project window (see Figure 5), select the slides and regions to be included in the analysis by checking the corresponding boxes and press Next.
Note: One can initiate a new TissueQuest analysis anytime with differently set parameters based on the same TissueFAXS acquisition; alternatively, one can append an already performed analysis with new selected regions from the same acquisition project and reapply the same analysis strategy onto those, for uniformity.
Figure 5. Screenshot of the “Choose TissueFAXS Project” dialog window
4. In the TissueQuest Project window, select the desired location to store the project analysis. By default, a project folder will be created in the same folder where the initial image acquisition project is stored. Optionally, you can add notes describing the experimental details. Click Next.
5. In the Choose Markers window (see Figure 6A), add/remove/rename the fluorescence markers (i.e., channels) for the desired analysis (e.g., DAPI and Texa). By default, DAPI is the master channel, as the single cells are identified based on the nucleus staining. You can change this if you use other nuclear dye.
6. Click on Add Dots Virtual Marker (see Figure 6B) to add a virtual channel on TxRed to define Texa_DOTS_ON_DAPI. Click OK.
Figure 6. Snapshot of the Choose Markers dialog window and the Add Dots Marker option. A. DAPI and Texa markers are shown by default based on the information from the scanned project. B. One can add additional markers, such as a dots marker using a virtual marker. C. The Texa_DOTS_ON_DAPI marker is added to the markers list of the project to analyze Alexa Fluor 594 labeled nuclear foci that overlaps with the Hoechst stained nucleus detected on DAPI.
7. Choose Various Shapes 2.0 and Select only events entirely in ROI for analysis and press Next (see Figure 6C).
8. In the Choose Measured Parameters window (see Figure 7), select the parameters to be used (e.g., DAPI Area, DAPI Mean intensity, DAPI Compactness, Texa Mean Intensity) for quantitative analysis on each channel. Additionally, choose Dots count as a parameter on the created Texa_DOTS_ON_DAPI virtual channel.
Note: For co-cultures where cell lines can be distinguished based on nuclei elongation, one can use either the eccentricity parameter [4] or the Feret ratio on DAPI [13] (both defining nuclei circularity index based on the ratio between their major and minor axes).
9. Add or remove specific scattergrams or histograms from the diagram list of the analysis project. Proceed to the analysis by clicking on the Finish button.
Note: The sequence of diagrams can also be changed at any step during analysis using the dedicated Diagrams action button.
Figure 7. Screenshot of the Choose Measured Parameters dialog window
10. When the analysis window opens (see Figure 8A), click each region one by one and wait for the cache to be built for all of them. Progress is visible by the filling of an orange status bar (see Figure 8B).
Figure 8. Screen capture of the TissueQuest analysis window (left screen) depicting menu tabs in the menu bar and a list of samples in the Project Browser mini-window. By default, an image of the first simple region in the sample list is open (A). Cache building progress is visible in the orange status bar and the date and time when the process is finished is shown at the end (B).
11. Select the region drawn onto the positive control sample (i.e., cells treated with CisPt to induce DNA damage response) from the sample list to define an area for positive signal threshold setup. Double-click on it to open the region in a separate Input window on the right-hand screen (see Figure 9A). The tab corresponding to the original image (i.e., merged channels color view) visualization mode is selected. Zoom in and use the rectangle Add Rectangular ROI tool to enclose a representative cell subpopulation with various levels of signals on the TxRed channel including the dimmest and the brightest foci, and assign it a name (e.g., “Set-up region”) in the sample list (see Figure 9B). Choose this sub-region when prompted by the software to use it for fine-tuning the signal thresholds for each parameter; it will appear on the right-side screen (see Figure 9C).
Note: If one used the entire sample region for analysis setup, it would take a considerable amount of time for each iteration step in the algorithm.
Figure 9. Screen captures of the software windows used to segment the nuclei and analyze the DAPI and TxRed signals. Key actionable buttons and tabs are visible. A. Left-side and right-side monitor views of the TissueQuest software. B. Creation of a setup ROI within the positive control sample. C. Fine-tuning of marker pixel intensity thresholds for image segmentation.
12. To identify single cells, first choose in the right-side screen Input window (see Figure 10A) the tabs corresponding to the DAPI channel to visualize nuclei in grayscale mode and Shades Overlay to observe the segmentation result in green. Then, fine-tune algorithm parameters in the DAPI tab of the Markers panel (see Figure 10B) and press the Analyze button marked with the Settings symbol above in the Input window. Do as many iterations as needed until the nuclei are correctly segmented (i.e., the encircled green masks overlap optimally with the visible DAPI-positive nuclei).
Note: We found the nuclei size of 40 (a.u.) and the DAPI threshold range of 12–80 to be optimal for our cells (see Figure 10B).
Note: Dots corresponding to individual cells analyzed will appear in the scattergrams. Their projection on the axes corresponds to the parameter value for the specific cell.
Figure 10. Obtaining segmentation masks. A. Magnified view of markers analysis criteria input panel. B. Example of a result obtained from the nuclei segmentation. C. Example of a result obtained from the Texa nuclear signal segmentation for p-γ-H2AX. D. Example of a result obtained from the Texa_DOTS_ON_DAPI nuclear p-γ-H2AX foci signal segmentation.
13. To identify the TxRed signal, choose “Texa” from the Input tab and from the Markers tab. Here, select “Yes” for “Use Nuclei Mask” in order to use the generated nuclei mask on DAPI as the location for Texa signal quantification (see Figure 10C). Then, fine-tune background threshold parameters to optimize signal detection.
Note: We found a background threshold of 5 to be optimal for our TxRed signal-to-noise ratio (see Figure 10C).
14. To identify the foci, first choose the Tab corresponding to “Texa DOTS_ON_DAPI” virtual channel to visualize nuclear foci encircled in yellow. Then, fine-tune algorithm parameters in the Texa tab of the Markers panel (see Figure 10D) and press the Analyze button. Do as many iterations as needed until the p-γ-H2AX foci are correctly segmented (i.e., the encircled yellow regions overlap optimally with the visible foci).
15. After the segmentation is done, set up the cutoff thresholds for each parameter analyzed (i.e., DAPI Area, DAPI Mean intensity, DAPI Compactness, DAPI Eccentricity, Texa Mean Intensity, Texa DOTS_ON_DAPI). First, make sure that all necessary scattergrams (2-parameter diagrams) and histograms (1-parameter diagrams) are visible. To add or remove elements, select the Manage Diagrams icon in the upper right Results menu above the scattergrams and histograms series and select the ones needed from the list. Press Close. To set up cell population thresholds, click on the Set Cutoff button in the upper left corner of each diagram and apply it with the left click of the mouse in the desired position.
Critical: Use forward and backward gating after each refining step to check if the negative and positive events (i.e., cells) are properly identified (see Figure 11). Right-click on the selected scattergram and select View Backward Data For Left Quadrants to visualize in red the events negative for the parameter detected on the horizontal axis (see Figure 11A for an example; Texa negative events in the second scattergram are shown in red). Right-click on the selected scattergram and select View Backward Data For Lower Quadrants to visualize in red the events negative for the parameter detected on the vertical axis. Right-click on the selected histogram and select View Backward Data For Left Quadrants to visualize in red the events negative for the parameter detected on the horizontal axis. Alternatively, the detected positive events can also be visualized by backward gating. Forward gating can be used by double-clicking to select a cell and check its quadrant location (see Figure 11B). After each step, check if false negative or false positive events are erroneously identified. If this is the case, adjust cutoff values by pulling it with the mouse toward the correct direction to include all positive events in the analysis and exclude all negative ones. Do this for all diagrams in the series.
Figure 11. Backward and forward gating tools. A. Use of backward gating to visualize the impact of choosing a certain cutoff value. B. Use of forward gating to visualize the signal intensity for a certain cell.
16. After the cell population cutoffs are set up, initiate the gating sequence. For this, select each scattergram in the series and generate gates to define the cell population to be analyzed and filter out all unwanted events. To draw gates, press one of the three gate shape icons depicted in blue for the specific scattergram (rectangle/circle/freeform), define the events population by left-clicking, and release it after you enclose the cell subset. You can readjust by pulling the shape sides. Gate the intact cells by selecting the upper-right quadrant of the DAPI Area/DAPI Mean Intensity scattergram. Go to the Manage Gates button and assign a name and color (e.g., blue) to gate the “single cells.” Then, go to the DAPI Compactness/DAPI Mean Intensity scattergram and select the previously gated events by choosing the “single cells” gate upon checking the Configure button in the upper-left corner. In this scattergram, define the “compact nuclei” gate to remove irregular artifacts from the analysis by selecting the events on the upper side. If you need to select a cell line based on nuclei elongation (as in Orobeti et al. [4]), draw two gates to define the “elongated/non-elongated nuclei” on the DAPI Eccentricity/DAPI Mean intensity scattergram after choosing the “compact nuclei” gate as input. Finally, use a histogram to depict “Texa DOTS_ON_DAPI” cells signal distribution on the selected cell subpopulation in the input gate (e.g., A375, NHEM, or co-culture). An example of a gating strategy is available in Figure 8 of the article Orobeti et al. [4].
Critical: Use backward gating after each refining step to check if the gated events are properly identified. Left-click the gate contour, right-click on it, and select View Backward Data For Gate. Do this for all diagrams in the series.
17. Analyze the entire region from which the setup region is a part. For this, click on the region name in the list of samples and then right-click Analyze from the options drop-down menu. When the analysis is finished, the region will appear as “Processed” in the sample list.
Note: Create exclusion gates for all areas containing artifacts in the sample before initiating analysis. For this, select Add Freedrawn Exclusion from the Freedrawn Tool icon on the left-side screen.
18. Visualize the results on the diagrams and refine cutoff values and gates, if necessary.
19. To batch process all the images, select each region to apply the same strategy and press Analyze. Repeat for all regions to be analyzed. Check if the events fit into the scattergrams and histograms for all samples and regions. Refine limits if necessary. Set the same axes maximum values and gate settings to all regions. For this, right-click on the diagram and select Set max value on: x, where x is the parameter displayed on the axis to be updated. To extend the settings to all analyzed regions, open the Manage Diagrams dialog window and select Apply Diagram Set to All Samples and their Regions. Refine gates if necessary, and batch apply the changes to all analyzed regions. Right-click on each gate and select Propagate gate x (where x is the gate name) and then All Samples and their Regions when prompted by the dialog window.
Data analysis
Data analysis consists of two main actions:
a. Exporting the statistics from the TissueQuest software as a report.
b. Exporting raw data in Excel spreadsheet(s) to be further analyzed in GraphPad Prism to generate graphs of the results, to calculate standard deviation, and evaluate the statistical significance of the possible observed differences between experimental conditions.
We will further indicate all necessary steps to perform the analysis of the image cytometry quantification for foci analysis, similarly to data presented in Orobeti et al. [4].
1. To export statistical data computed by TissueQuest, select Tools/Statistics Report from the menu bar.
2. In the open Statistics Report window, select the input regions from the sample list and perform the following steps to export for each sample the events count (e.g., total, A375 only, and NHEM only), p-γ-H2AX mean fluorescence intensity, or percentage of p-γ-H2AX-positive cells for all cell populations of interest:
a. Choose New Column/Global Measurements Parameters, leave Events Count under the Column Name, and select Events Count under Global Measurement.
b. For quantifying p-γ-H2AX mean fluorescence intensity (MFI), choose New Column/Predefined, pgH2AX MFI under Column Name, and select predefined value unit Mean of Mean Intensity of Texa marker from the propagated gate named compact nuclei.
Note: To analyze only a cell line from the co-culture, choose elongated/non-elongated under Select propagated gate.
c. For quantifying the percentage of p-γ-H2AX+ cells, choose New Column/Predefined, %pgH2AX+ cells under Column Name, and select predefined value unit Percent from the propagated gate named “pgH2AX+”.
3. To export raw data for analysis into GraphPad, select each analyzed sample/region and click the Spreadsheet icon in the upper-right Results window next to Diagram Options and Manage Gates icons. In the open Raw Data table (see Figure 12A), filter events sequentially (using thresholds previously established in step F15) based on DAPI compactness (see Figure 12B) and DAPI eccentricity. Then, export list of the currently selected region as an Excel file (press Export To Excel) to extract the Texa_DOTS_ON_DAPI-Dots Count values to import into GraphPad (see next step).
Figure 12. Exporting numeric values associated with the analyzed events. A. Screenshot of TissueQuest Raw Data table associated with one selected region in the sample list. B. Command to filter the events using the cutoff value. An example is given for the DAPI compactness parameter.
4. Save the Excel file with the name of the cell line and the quantified parameter.
5. Repeat steps 3–4 to export data for all analyzed samples/regions.
6. Open GraphPad to initiate the comparative analysis of foci count for all conditions tested within the experiment. Select the Column graph type option (see Figure 13) that allows comparison between different (radiation) treatment conditions from the same irradiation session for “number of p-γ-H2AX foci per nucleus.”
Note: Expertise using GraphPad Prism is required to obtain the described graphs and statistical data analysis. For this, consult Prism User Guide in the Help menu tab and Prism Academy online at https://www.graphpad.com/prism-academy.
Figure 13. Screen capture of GraphPad dialog window. Selection of the Column graph template.
7. Transfer data from Excel to GraphPad. For this, rename “Data 1” table with the name of your experimental comparison. Assign names to column titles (e.g., non-irradiated, X-ray-irradiated, etc.). Copy and paste data from each sample spreadsheet into the specific column in each table to generate graphs for co-culture (e.g., A375&NHEM) and single cultures (e.g., A375, NHEM).
8. On the autogenerated graph, double-click to refine the format (see Figure 14).
Figure 14. Screen capture of the Format Graph dialog window. Graph formatting options are visible.
9. To evaluate potential differences in double-strand DNA breaks induction, perform statistical comparisons using a Mann–Whitney test. A p-value < 0.05 is considered to indicate statistical significance. For this, click Analyze in the Analysis tab of the menu bar. Then, choose t tests from the Column Analysis stack, select the samples for which you are doing comparative analysis, and click OK. Choose a two-tailed unpaired t-test without assuming Gaussian distribution followed by a Mann-Whitney test to compare ranks.
Note: If the data is parametric, a Student’s t-test may be used instead of the Mann-Whitney test. A Shapiro-Wilk test can be used in GraphPad to determine whether a data set is parametric or non-parametric.
10. Represent the adjusted p-values on the graph for comparisons of each radiation condition with the non-irradiated control as star symbols. For this, choose Draw from the menu bar and select either Manually Add Lines With Text to select a format or Choose Pairwise Comparisons to plot for automatic representation of the statistical results.
11. Export the figures separately or as layout. Images should have at least 300 dpi (600 dpi for colored graphs, RGB quality) and be exported as .tiff for optimal publication quality resolution.
Note: Researchers without access to TissueQuest software can use other accessible image analysis software for foci count determination [11,12]. For this, export the “dots per nucleus” data according to the specific tool chosen and use it for statistical analysis in GraphPad (steps 6–11 presented above) or Excel.
Validation of protocol
This protocol has been used and validated in the following research article(s):
• Orobeti et al. [4]. First in vitro cell co‐culture experiments using laser‐induced high‐energy electron FLASH irradiation for the development of anti‐cancer therapeutic strategies. Scientific Reports (Figure 2, Figure 3, Figure 4, Figure 5, Figure 6, Figure 8).
General notes and troubleshooting
General notes
1. For more detailed indications on the use of TissueFAXS and TissueQuest software packages, please consult the user guides in the specific software menu for updated information specific to the versions available on your system or the respective printed versions received at installation.
Troubleshooting
Problem 1: Samples present artifacts.
Possible cause: Coverslips or slides were not cleaned; buffers contained aggregates; use enough mounting to prevent sample drying and air bubbles formation.
Solution: A) (wet lab procedures) Pass the coverslips through MilliQ ultrapure water using fine forceps before mounting on the slide to eliminate dust particles and aggregates and to prevent PBS crystals from forming. Clean the slide with 70% ethanol using Kimwipes. Filter buffer solutions used for immunofluorescence staining. B) (software procedure) Draw exclusion areas around the artifacts to remove them from the analysis. Any area with a different background level within the sample induces threshold errors in the analysis, thereby affecting the quality of the quantitative analysis results. Therefore, it should be removed.
Problem 2: Nuclei segmentation results are not precise.
Possible cause: Signal-to-noise ratio is deficient; DAPI analysis parameters are not optimal.
Solution: A) (wet lab procedures) Prevent background by carefully and thoroughly washing the samples after each staining step and by not using excessive mounting solution. B) (software procedure) Also, carefully optimize the exposure time before sample scanning to obtain a bright signal without increasing background levels. To optimize nuclei masks during segmentation, fine-tune background threshold parameters until the masks overlap with the desired signal. To observe background gray levels, hover the mouse over the space between nuclei, and the value will be shown in the upper-left corner of the image. The lower signal threshold level should be above that of the background. Finally, if two nuclei are seen as one, a line can be used to manually separate them to optimize correct segmentation; alternatively, exclude doublets and aggregates during analysis when applying the first gate using DAPI Area measurement.
Acknowledgments
This research was supported by IFA (Institute of Atomic Physics) through the ELI-RO_2020_11 Project, No. 01/2020 and ELI-RO_2024_10 Project, No. 10/2024; and the Romanian Ministry of Education and Research under the Romanian National Nucleus Program LAPLAS VII under Contract No. 30N/2023. This work received funding from the European Union’s Horizon 2020 Research and Innovation Program under Grant Agreement No. 871124 Laserlab-Europe. S.O. and L.E.S. acknowledge the partial support of Project No. 1/2023 of the Research Program of the Institute of Biochemistry of the Romanian Academy. S.O., I.D., A.B., I.T., and F.S. acknowledge the support of National Interest Infrastructure facility IOSIN – CETAL at INFLPR. We acknowledge the authors of the article entitled “First in vitro cell co‐culture experiments using laser‐induced high‐energy electron FLASH irradiation for the development of anti‐cancer therapeutic strategies” in which the use of this protocol was first described [4].
Competing interests
The authors declare no competing interests.
References
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Fuchs, T., Szymanowski, H., Oelfke, U., Glinec, Y., Rechatin, C., Faure, J. and Malka, V. (2009). Treatment planning for laser-accelerated very-high energy electrons. Phys Med Biol. 54(11): 3315–3328.
Orobeti, S., Sima, L. E., Porosnicu, I., Diplasu, C., Giubega, G., Cojocaru, G., Ungureanu, R., Dobrea, C., Serbanescu, M., Mihalcea, A., et al. (2024). First in vitro cell co-culture experiments using laser-induced high-energy electron FLASH irradiation for the development of anti-cancer therapeutic strategies. Sci Rep. 14(1): 14866.
Belyaev, I. Y. (2010). Radiation-induced DNA repair foci: spatio-temporal aspects of formation, application for assessment of radiosensitivity and biological dosimetry. Mutat Res. 704(1-3): 132–141.
Vignard, J., Mirey, G. and Salles, B. (2013). Ionizing-radiation induced DNA double-strand breaks: a direct and indirect lighting up. Radiother Oncol. 108(3): 362–369.
Wang, J. Y. J. (2019). Cell Death Response to DNA Damage. Yale J Biol Med. 92(4): 771–779
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Vasireddy, R. S., Sprung, C. N., Cempaka, N. L., Chao, M. and McKay, M. J. (2010). H2AX phosphorylation screen of cells from radiosensitive cancer patients reveals a novel DNA double-strand break repair cellular phenotype. Br J Cancer. 102(10): 1511–1518.
Cucinotta, F. A., Pluth, J. M., Anderson, J. A., Harper, J. V. and O'Neill, P. (2008). Biochemical kinetics model of DSB repair and induction of gamma-H2AX foci by non-homologous end joining. Radiat Res. 169(2): 214–222.
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Article Information
Publication history
Received: Oct 14, 2024
Accepted: Dec 23, 2024
Available online: Jan 19, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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5,209 | https://bio-protocol.org/en/bpdetail?id=5209&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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Peer-reviewed
Versatile Click Chemistry-based Approaches to Illuminate DNA and RNA G-Quadruplexes in Human Cells
AP Angélique Pipier
DM David Monchaud
Published: Vol 15, Iss 3, Feb 5, 2025
DOI: 10.21769/BioProtoc.5209 Views: 201
Reviewed by: Emilie Besnard Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in RSC Chemical Biology May 2023
Abstract
The existence and functional relevance of DNA and RNA G-quadruplexes (G4s) in human cells is now beyond debate, but how did we reach such a level of confidence? Thanks to a panoply of molecular tools and techniques that are now routinely implemented in wet labs. Among them, G4 imaging ranks high because of its reliability and practical convenience, which now makes cellular G4 detection quick and easy; also, because this technique is sensitive and responsive to any G4 modulations in cells, which thus allows gaining precious insights into G4 biology. Herein, we briefly explain what a G4 is and how they can be visualized in human cells; then, we present the strategy we have been developing for several years now for in situ click G4 imaging, which relies on the use of biomimetic G4 ligands referred to as TASQs (for template-assembled synthetic G-quartets) and is far more straightforward and modular than classically used immunodetection methods. We thus show why and how to illuminate G4s with TASQs and provide a detailed, step-by-step methodology (including the preparation of the materials, the methodology per se, and a series of notes to address any possible pitfalls that may arise during the experiments) to make G4 imaging ever easier to operate.
Key features
• MultiTASQs are clickable probes usable for the detection of cellular DNA and RNA G-quadruplex (G4).
• In situ click chemistry relies on the labeling of G4 by clickable probes once in their cellular binding sites.
• Experiments can be performed by incubating the clickable probe either in live cells or in fixed cells.
• The published but unoptimized protocol is now totally revised to allow for reliable G4 detection in human cells.
Keywords: DNA/RNA G-quadruplex (G4) Optical imaging Click chemistry Template-assembled synthetic G-quartets (TASQs) Clickable MultiTASQs
Graphical overview
Background
Guanine (G)-rich DNA and RNA sequences are being studied for their ability to fold into G-quadruplex (G4) structures [1–3]. G4-prone sequences are widespread in our genome and transcriptome; it has now been established that G4s are key players in virtually all genomic transactions, from the regulation of genetic and epigenetic events to chromatin organization [1–8]. G4-prone sequences have been identified in silico by various methods [9], which showed that >1 million sequences can fold into G4s in our genome [10] and transcriptome [11] (referred to as G4s and rG4s for DNA and RNA G4s, respectively). This formation has been demonstrated at both genome-wide and transcriptome-wide scales using methods being classified as in vitro (that is, using purified DNA and RNA samples: G4-seq [12] and G4DP-seq [13] for G4s, rG4-seq [14] for rG4s) or in vivo (that is, in a functional cellular context: G4 ChIP-seq [15], G4 CUT&Tag [16], and G4access [17] for G4s, G4RP-seq [18] for rG4s). These sequencing-based methods confirmed the prevalence of G4 landscapes in human cells, with the identification of hundreds of thousands in vitro G4s and tenths of thousands in vivo G4s [19].
The cellular roles of G4s are beyond the scope of the present article; interested readers are invited to consult recent reviews on this topic (for G4s, see [1,2,7]; for rG4s, see [3–6,8]). We would like to report here on a convergent strategy we have been developing for several years now, based on the use of smart molecular tools named template-assembled synthetic G-quartets (TASQs) (Figure 1) [20]. Our aims are to i) select the best-suited TASQs among those now available as a function of the intended application and ii) use this tool for addressing more than one chemical biology challenge (optical imaging, affinity purification, chemoproteomics, etc.). TASQs are structurally dynamic molecules comprising a central template surrounded by four flexible arms ending with a G unit; they are thus biomimetic ligands that stabilize G4s by interacting between two G-quartets, one from the G4, and one from the TASQ. We developed the smart PNADOTASQ [21], twice-as-smart PyroTASQ [22] and NaphthoTASQ (or N-TASQ) [23], biotinylated BioTASQ [24], BioCyTASQ [25] and BioTriazoTASQ [26], and multivalent MultiTASQ and azMultiTASQ [27]. These TASQs fold into their G4-affinic closed conformation only upon interaction with G4s (that is, only when a G-rich sequence folds into a G4 structure), which makes them uniquely actively selective for their G4 targets [20,21].
Figure 1. Schematic representation of the folding of a G-quadruplex (G4) structure from a guanine (G)-rich DNA sequence. Chemical structure of TASQs, either MultiTASQ or azMultiTASQ, and of their derivatives after click chemistry (CuAAC or SPAAC)-based coupling with a fluorophore; the resulting conjugate aimed at being used to visualize G4s in human cells.
Here, we report on the use of MultiTASQs to illuminate G4s, given that MultiTASQ and azMultiTASQ are the sole clickable TASQs from this family of compounds. Several strategies have been pursued to visualize G4s in cells [28–30], but we are particularly interested in a quite efficient two-step methodology based on the fluorescent tagging of chemical ligands once in their genomic binding site by in situ click chemistry [31]. This yet indirect approach allows for better control of the fluorescence readout, which makes it more versatile. This approach, pioneered in the G4 field by Rodriguez et al. in 2012 [32], is here extended using now commercially available clickable TASQs. We thus provide all necessary technical details for performing in situ click G4 imaging using MultiTASQs, focusing on the step-by-step methodology and all possible cell manipulations to increase the quality of the collected images and all necessary controls to exploit the obtained results in a reliable manner.
Materials and reagents
Note: Caution must be exercised when performing TASQs staining experiments; the most important features are gathered in the different notes below (see General notes 1–4), which must be read before preparing the materials needed for the experiments. All solutions must be prepared (unless otherwise stated) in ultrapure water (resistivity ≥ 18 MΩ·cm at 25 °C), and molecular biology and/or analytical grade reagents must be used. Solutions can be prepared and stored at room temperature (unless otherwise stated); some of them need to be prepared under a sterile environment in the biological safety cabinet (BSC) and/or to be filtered (PES 0.22 μm bottle-top vacuum filter system or syringe filter system, depending on volume).
Biological materials
1. HeLa cell line (ATCC, catalog number: CCL-2)
Reagents
Note: As not all reagents and/or solutions are absolutely required (see General notes 1–4) and some are mutually exclusive, we strongly recommend reading the procedure in its entirety to determine if they should be used.
Note: We provide the list of reagents we used, but reagents from alternative suppliers are acceptable as long as they are molecular biology and/or analytical-grade reagents.
1. Dulbecco’s modified Eagle medium (DMEM) containing sodium pyruvate, high glucose, and GlutaMAX (Gibco, catalog number: 31966021); store at 4 °C
2. Fetal bovine serum (FBS) 100% (Dutscher, catalog number: on request); store at -80 °C
3. Penicillin/streptomycin 10,000 U/mL (Gibco, catalog number: 15140122); store at -20 °C
4. Trypsin-EDTA (0.25%), phenol red (Gibco, catalog number: 25200056); once thawed, store at 4 °C
5. PBS 10× (Eurobio, catalog number: ET330A)
6. Pyridostatin hydrochloride (PDS) (Sigma-Aldrich, catalog number: SML2690-5MG)
7. Dulbecco’s PBS 10× (DPBS) (Dutscher, catalog number: X0515-500)
8. TBS 10× (Euromedex, catalog number: ET220-B)
9. Triton X-100 (Sigma-Aldrich, catalog number: X100-100ML); store at 4 °C
10. Tween-20 (Fisher BioReagents, catalog number: BP337-500)
11. 1,4-Piperazinediethanesulfonic acid (PIPES) (Sigma-Aldrich, catalog number: P8203)
12. Sodium chloride (NaCl) (Sigma-Aldrich, catalog number: 71376-1KG)
13. Sucrose (Sigma-Aldrich, catalog number: S0389)
14. Magnesium chloride (MgCl2) (Sigma-Aldrich, catalog number: M8266)
15. RNase A 10 mg/mL (Thermo Scientific, catalog number: EN0531); store at -20 °C
16. TURBOTM DNase kit 2 U/μL containing TURBOTM DNase and 10× TURBOTM DNase buffer (Invitrogen, catalog number: AM2238); store at -20 °C
17. Methanol (MeOH) (Carlo Erba Reagents, catalog number: 412383); store at -20 °C
18. Paraformaldehyde (PFA) 16% w/v aqueous solution, methanol free (Thermo Scientific, catalog number: 28908); once opened, store in a 15 mL Falcon tube at 4 °C
19. Bovine serum albumin (BSA) (Seqens, catalog number: 1000-70); store at 4 °C
20. Fish gelatin (Sigma-Aldrich, catalog number: G7041); store at 4 °C
21. MultiTASQ (Merck, catalog number: SCT247); store at -20 °C
22. azMultiTASQ (not commercially available, contact D. Monchaud); store at -20 °C
23. Alexa FluorTM 594 azide (AF-az) 1 mM in DMSO [Alexa FluorTM 594 carboxamido-(6-azidohexanyl), triethylammonium salt], mixed isomers (Invitrogen, catalog number: A10270); store at -20 °C, avoid light
24. Click-iTTM Alexa FluorTM 594 sDIBO alkyne (DIBO-AF) 5 mM in DMSO (Invitrogen, catalog number: C10407); store at -20 °C, avoid light
25. Copper(II) sulfate (CuSO4) (Sigma-Aldrich, catalog number: C7631-250G)
26. IGEPAL CA-630 (Sigma-Aldrich, catalog number: I8896)
27. Sodium ascorbate (NaAsc) (Sigma-Aldrich, catalog number: 11140)
28. 4',6-Diamidino-2-phenylindole dihydrochloride (DAPI) (Sigma-Aldrich, catalog number: D9542)
29. VECTASHIELD® antifade mounting medium (VectorLab, catalog number: H1900); store at 4 °C
30. Transparent nail polish
Solutions
Note: Filtered solutions must be aliquoted to avoid cross-contamination and/or bacterial counterstaining with DNA intercalating dyes. Solutions with adjusted pH must be prepared with a sufficient volume of ultrapure water (ca. 90% of final volume) to resuspend powder by mixing, then pH must be adjusted, and finally, the solution can be completed with ultrapure water to final volume.
1. PDS 20 mM in DMSO
Note: Please refer to the manufacturer’s certificate of analysis to obtain the molecular weight of PDS with the number (y) of molecules of HCl as counterion and with the number (z) of molecules of water [final molecular weight (g/mol) = 596.64 + y × 36.46 + z × 18.02]. Store at -80 °C.
2. MultiTASQ or azMultiTASQ 10 mM in nuclease-free water
Note: Molecular weight of MultiTASQ: 1801.12 g/mol. Molecular weight of azMultiTASQ: 1846.16 g/mol. To avoid batch-to-batch variations in concentrations of stock solution of TASQ, we recommend measuring the absorbance of TASQ’s guanines at λ = 246 nm (see section A. Preparation of TASQ). Store at -20 °C.
3. AF-az 1 mM in DMSO
Note: Please refer to the manufacturer’s molecular weight depending on the number of triethylammonium molecules as counterion (molecular weight should be around 1,050 g/mol for 2 counterions). Store at -20 °C, avoid light.
4. DIBO-AF 5 mM in DMSO
Note: Please refer to the manufacturer’s molecular weight depending on the number of triethylammonium molecules as counterion (molecular weight should be around 1,200 g/mol for 2 counterions). Store at -20 °C, avoid light. Intermediate dilution of 0.5 mM of DIBO-AF in water (10% final concentration of DMSO) is more handful for SPAAC mix.
5. DAPI 1 mg/mL in water
Note: Store at 4 °C, avoid light.
6. Sterile PBS 1× (see Recipes)
7. Sterile supplemented DMEM (see Recipes)
8. Filtered DPBS 1× (see Recipes)
9. Filtered DPBS-Tx (see Recipes)
10. Filtered TBS-Tw (see Recipes)
11. PIPES 100 mM, pH 7.0 (see Recipes)
12. NaCl 1 M (see Recipes)
13. MgCl2 1 M (see Recipes)
14. Filtered sucrose 1 M (see Recipes)
15. Filtered cytoskeletal pre-extraction (CSK) buffer (see Recipes)
16. Filtered BSA blocking buffer (see Recipes)
17. Filtered FBS blocking buffer (see Recipes)
18. MultiTASQ 20 μM (200 μL) for post-fixation staining (see Recipes)
19. azMultiTASQ 20 μM (200 μL) for post-fixation staining (see Recipes)
20. CuSO4 100 mM (see Recipes)
21. IGEPAL CA-630 10% (see Recipes)
22. NaAsc 300 mg/mL (150×) (see Recipes)
23. CuAAC mix (see Recipes)
24. SPAAC mix (see Recipes)
Recipes
1. Sterile PBS 1× (500 mL)
Note: Prepare for cell culture (sterilize by filtration or autoclaving) and open only under sterile conditions.
Reagent Final concentration Quantity or Volume
PBS (10×) 1× 50 mL
H2O n/a 450 mL
Total n/a 500 mL
2. Sterile supplemented DMEM (555 mL)
Note: Prepare directly in a DMEM bottle (from the manufacturer) and open only under sterile conditions.
Reagent Final concentration Quantity or Volume
DMEM (1×) n/a 500 mL
FBS (100%) 10% 55 mL
Penicillin/Streptomycin (100%) 1% 5.5 mL
Total n/a ca. 555 mL
3. Filtered DPBS 1× (500 mL)
Note: Filter and aliquot 50 mL for use. Keep a 50 mL aliquot at 4 °C as cold DPBS.
Reagent Final concentration Quantity or Volume
DPBS (10×) 1× 50 mL
H2O n/a 450 mL
Total n/a 500 mL
4. Filtered DPBS-Tx (500 mL)
Note: Filter and aliquot 50 mL for use.
Reagent Final concentration Quantity or Volume
DPBS (10×) 1× 50 mL
H2O n/a 450 mL
Triton X-100 (100%) 0.1% 500 μL*
Total n/a ca. 500 mL
*Tip needs to be cut due to Triton X-100 viscosity.
5. Filtered TBS-Tw (500 mL)
Note: Filter and aliquot 50 mL for use.
Reagent Final concentration Quantity or Volume
TBS (10×) 1× 50 mL
H2O n/a 450 mL
Tween-20 (100%) 0.1% 500 μL*
Total n/a ca. 500 mL
*Tip needs to be cut due to Tween-20 viscosity.
6. PIPES 100 mM, pH 7.0 (50 mL)
Note: Store at 4 °C up to a year. We recommend controlling pH if PIPES has not been used in the past 3 months.
Reagent Final concentration Quantity or Volume
PIPES 100 mM 1.51 g
H2O n/a n/a*
NaOH 10 M n/a Until reaching pH 7.0**
Total n/a 50 mL
*Add 40 mL of ultrapure water before adjusting pH. Once pH is adjusted, complete with water to obtain 50 mL of solution.
**pH equilibration with NaOH 10 M is necessary to dissolve PIPES; adjust pH to 7.0 by adding dropwise NaOH in solution under stirring.
7. NaCl 1 M (50 mL)
Note: Store at 4 °C for up to a year.
Reagent Final concentration Quantity or Volume
NaCl 1 M 2.92 g
H2O n/a n/a*
Total n/a 50 mL
*Add 40 mL of ultrapure water and allow the solution to mix completely before adding any more water.
8. MgCl2 1 M (50 mL)
Note: Store at 4 °C for up to a year, avoid light.
Reagent Final concentration Quantity or Volume
MgCl2 1 M 4.76 g
H2O n/a n/a*
Total n/a 50 mL
*Add 40 mL of ultrapure water and allow the solution to mix completely before adding any more water.
9. Filtered sucrose 1 M (50 mL)
Note: Filter and store at -20 °C to avoid contamination.
Reagent Final concentration Quantity or Volume
Sucrose 1 M 17.1 g
H2O n/a n/a*
Total n/a 50 mL
*Add 30 mL of ultrapure water and allow the solution to mix completely before adding any more water as sucrose will take a substantial volume when fully dissolved.
10. Filtered cytoskeletal pre-extraction (CSK) buffer (50 mL)
Note: Filter with a 0.22 μm PES filter for a syringe and aliquot 5–10 mL of CSK in vials under sterile conditions. Store aliquoted vials at -20 °C. Avoid freeze/thaw cycles of aliquoted vials (2–3 cycles maximum).
Reagent Final concentration Quantity or Volume
PIPES (100 mM, pH 7.0) 10 mM 5 mL
NaCl (1 M) 100 mM 5 mL
Sucrose (1 M) 300 mM 15 mL
MgCl2 (1 M) 3 mM 150 μL
H2O n/a 24.5 mL
Triton X-100 (100 %) 0.7% 350 μL*
Total n/a 50 mL
*Tip needs to be cut due to Triton X-100 viscosity. Homogenize before adding Triton X-100 to avoid foaming.
11. Filtered BSA blocking buffer (500 mL)
Note: We found that the appropriate blocking buffer for MultiTASQ is BSA blocking buffer, and that for azMultiTASQ is FBS blocking buffer (see Troubleshooting 4; see Figure 10).
Filter and aliquot 50 mL of blocking buffer in vials under sterile conditions. Store aliquoted vials at -20 °C. Once a vial is thawed, store at 4 °C for 2–3 months maximum.
Reagent Final concentration Quantity or Volume
BSA 1% (w/v) 5 g
Fish gelatin 0.2% (w/v) 1 g
DPBS (10×) 1× 50 mL
H2O n/a n/a
Triton X-100 (100 %) 0.1% 500 μL*
Total n/a ca. 500 mL
*Add 300 mL of water, allow BSA and fish gelatin to dissolve, and complete to 500 mL before addition of Triton X-100. The tip needs to be cut due to Triton X-100 viscosity. Homogenize before adding Triton X-100 to avoid foaming.
12. Filtered FBS blocking buffer (50 mL)
Note. We found that the appropriate blocking buffer for MultiTASQ is BSA blocking buffer, and that for azMultiTASQ is FBS blocking buffer (see Troubleshooting 4; see Figure 10).
Prepare and filter under sterile conditions with a 0.22 μm PES filter for a syringe. Store at 4 °C for 1–2 months maximum.
Reagent Final concentration Quantity or Volume
FBS (100 %) 2% 1 mL
DPBS-Tx n/a 49 mL
Total n/a 50 mL
13. MultiTASQ 20 μM (200 μL) for post-fixation staining
Note: Prepare just before incubation with MultiTASQ (see Procedure for post-fixation staining). This volume corresponds to the volume required for a well (see General note 5). This solution cannot be stored.
Reagent Final concentration Quantity or Volume
MultiTASQ (2 mM, see section A) 20 μM 2 μL
Filtered BSA blocking buffer n/a 198 μL
Total n/a 200 μL
14. azMultiTASQ 20 μM (200 μL) for post-fixation staining
Note: Prepare just before incubation with azMultiTASQ (see Procedure for post-fixation staining). This volume corresponds to the volume required for a well (see General note 5). This solution cannot be stored.
Reagent Final concentration Quantity or Volume
azMultiTASQ (2 mM, see section A) 20 μM 2 μL
Filtered FBS blocking buffer n/a 198 μL
Total n/a 200 μL
15. CuSO4 100 mM (1 mL)
Note: Store at 4 °C and avoid light.
Reagent Final concentration Quantity or Volume
CuSO4 100 mM 24.97 mg
H2O n/a 1 mL
Total n/a ca. 1 mL
16. IGEPAL CA-630 10% (1 mL)
Note: Store at 4 °C.
Reagent Final concentration Quantity or Volume
IGEPAL CA-630 (100%) 10 % 100 μL*
H2O n/a 900 μL
Total n/a 1 mL
*Tip needs to be cut due to IGEPAL CA-630 viscosity.
17. NaAsc 300 mg/mL (150×) (300 μL)
Note: Aliquot vials of 30 μL and store at -20 °C. Avoid freeze/thaw cycle as NaAsc is sensitive to oxidation.
Reagent Final concentration Quantity or Volume
NaAsc 300 mg/mL 90 mg
H2O n/a 300 μL
Total n/a ca. 300 μL
18. CuAAC mix (200 μL)
Note: Prepare just before “click” chemistry step with MultiTASQ (see Procedure). This volume corresponds to the volume required for a well (see General note 5). Add reagents in the order listed below. This solution cannot be stored.
NaAsc 30 mg/mL (or 15×) should be prepared in nuclease-free ultra-pure water before CuAAC mix; for 15 μL, add 1.5 μL of NaAsc 300 mg/mL to 13.5 μL of nuclease-free ultra-pure water.
Reagent Final concentration Quantity or Volume
PBS (1×) n/a 184 μL
IGEPAL (10%) 0.05% 1 μL
CuSO4 (100 mM) 1 mM 2 μL
AF-az (1 mM) 1 μM 0.2 μL
NaAsc (30 mg/mL or 15×) 2 mg/mL or 1× 13.3 μL
Total n/a ca. 200 μL
19. SPAAC mix (200 μL)
Note: Prepare just before “click” chemistry step with azMultiTASQ (see Procedure). This volume corresponds to the volume required for a well (see General note 5). This solution cannot be stored.
Reagent Final concentration Quantity or Volume
DIBO-AF (0.5 mM) 0.5 μM 0.2 μL
FBS blocking buffer (1×) n/a 200 μL
Total n/a ca. 200 μL
Laboratory supplies
1. PES filter 0.22 μm Stericup system (Sigma-Aldrich, catalog number: S2GPU10RE)
2. OmnifixTM solo 50 mL single-use syringe (Braun, catalog number: 4613503F)
3. PES filter 0.2 μm (MinisartTM high flow) for syringe (Sartorius, catalog number: 16532----------K)
4. Tissue culture treated flask (75 cm2) (Falcon, catalog number: 353136)
5. Glass coverslips 13 mm, #1.5 (VWR, catalog number: 631-0150)
6. Microscope glass slides 26 × 76 × 1 mm (ISO8037/I) (Epredia, catalog number: AA00000112E01MNZ10)
7. 24-well polystyrene microplate (2 cm2) (Falcon, catalog number: 353047) or 4-well polystyrene microplate (2 cm2) (SPL Life Science, catalog number: 30004)
Equipment
1. pH meter
2. Spectrophotometer (UV5Nano)
3. Biological safety cabinet
4. 37 °C, 5% CO2 incubator
Note: Equipment is dedicated to mammalian cell culture to avoid any bacterial contamination.
5. Aspiration system
Note: The aspiration system allows hazardous biological fluids (cell debris, fixative reagent, DAPI, etc.) to be disposed of.
6. Counting chamber (Malassez pattern)
7. Phase-contrast or brightfield microscope
8. Widefield microscope (Olympus, model: IX83)
a. DAPI filter set (Chroma, catalog number 49000)
b. TexasRed filter set (Chroma, catalog number 49306)
c. Optional for multiplexing: GFP filter set (Chroma, catalog number 49002) and/or Cy5 filter set (Chroma, catalog number 49009) and/or Cy7 filter set (Chroma, catalog number 49007)
Software and datasets
1. CellSens Dimension (Olympus, v3.2)
2. FIJI (ImageJ, v2.14.0)
3. Excel (Microsoft, v16.77.1)
4. Prism (GraphPad, v10.3.1)
5. Code has been deposited to GitHub (see Data analysis)
Procedure
Note: The described protocol is optimized for HeLa cells seeded in a 24-well plate and treated 24 h after seeding. For this type of protocol, be fast (but not furious): do not allow cells to dry but do not rush to prevent them from detaching.
A. Preparation of TASQs
1. After resuspension of TASQ (MultiTASQ and/or azMultiTASQ) at 10 mM in nuclease-free ultra-pure water, measure the absorbance of TASQ’s guanines with a spectrophotometer at λ = 246 nm.
a. Concentration is calculated as follows:
c
=
A
246
n
m
4
×
10700
×
l
, taking into account the optical path length of the spectrophotometer used (l) and the molar absorption coefficient corresponding to four guanines (4 × 10,700), according to Beer-Lambert formula.
Caution: Determination of blank value has to be done with the same water as the one used to resuspend TASQ.
Caution: Serial dilutions might be needed in particular if the absorbance values are out of the linear range of the spectrophotometer used and/or for a better correction of concentration.
Critical: We would like to stress that this molar extinction coefficient value is an approximation and, therefore, the corrected concentration does not represent absolute concentration but a standard for obtaining reproducible experiments from different production batches as the number of TFA molecules can vary (one molecule of TFA representing ca. 6.2% of TASQs molecular weight).
2. Adjust the concentration value of the TASQ stock solution.
3. Prepare a 2 mM dilution of TASQ in nuclease-free ultra-pure water.
Critical: Amines from guanines of TASQ can be degraded with time and repeated freeze/thaw cycles.
Note: Prepare aliquots with appropriate volume for staining experiments (2–5 μL of TASQs are needed per well, depending on live-cell or post-fixation staining). Keep at -20 °C for up to a year.
Note: If TASQ staining starts to fade and/or be unexpected, measure absorbance at λ = 246 nm to be sure of the integrity and concentration of the stock solution and prepare a new 2 mM dilution.
B. Cell seeding
1. Grow cells to reach ca. 70%–75% confluency on the day of seeding.
Note: A high confluency could impair doubling time and therefore processes (such as replication) allowing G4 detection.
2. Prepare sterile coverslips.
Note: A quick and efficient way to sterilize coverslips is to microwave them (dry, in a glass Petri dish) with a beaker of water aside for 5–10 min at maximal power.
3. Put one sterile coverslip per well in a 24-well plate, add 500 μL of supplemented DMEM per well, and incubate for at least 10 min at 37 °C.
Caution: To keep the coverslips sterile, we recommend using a sterile Pasteur or aspiration pipette to aspirate it and pinch the pipe to release it within the well.
Note: We observed that incubation of plates and coverslips with cell culture medium avoids central “halo-like” repartition of cells in the well, which ensures better conditions for treatments and imaging.
4. After trypsin-mediated harvesting of cells (performed according to standard protocols), count cells with a counting chamber.
Caution: Avoid leaving the cells for too long in trypsin alone as it could impact cell viability.
Note: Dilution of trypsin with supplemented DMEM is sufficient to abolish the deleterious impact of trypsin onto HeLa cells; no further centrifugation is required.
5. Prepare a solution of cells diluted in supplemented DMEM at 200,000 cells/mL.
Critical: We highly recommend optimizing the number of cells per well before using TASQ (see Troubleshooting 1 and 2). Indeed, several features of the cell line (including morphology, size, and doubling time) can influence the final confluency before the pre-fixation or fixation step. We observed for some cell lines that a high confluency could be deleterious for CSK pre-extraction, which leads to the loss of a high number of cells.
6. Add 500 μL/well of solution of diluted cells to obtain 100,000 cells/well corresponding to ca. 70% confluency after 24 h of incubation at 37 °C.
Note: We recommend treating after 24–48 h (that is, when cells recover their normal morphology) and monitoring the confluency during long live treatments (>4 h).
7. After 24 h, replace the medium with 500 μL of fresh medium. At this step, you can treat cells with drugs [e.g., Pyridostatin (PDS) [32,33] 10 μM for 4 h] and use TASQ as a post-fixation G4 probe (Figure 2A, see Procedure section D) or as a G4 ligand for live-cell experiments (Figure 2B, see Procedure section E).
Critical: We highly recommend using a well-established G4 ligand (such as PDS) as a positive control; PDS is herein used to stabilize cellular G4s and thus to increase the number of G4 foci (referred to as positive control herein; see the Validation section below), which is important notably during the optimization of TASQs staining.
Note: If you plan to treat several wells, see General note 5.
Figure 2. Workflow of (A) post-fixation or (B) live-cell TASQs staining. *: These steps (pre-extraction and/or nuclease treatment) are optional. **: Fixation duration is based on longest fixation (i.e., 15 min for PFA fixation vs. 5 min for methanol fixation). ***: When extensive washing is not required, the washing time is fixed at 5 min (please refer to the protocol for washing buffer used).
C. Preparation of nucleases treatment (if needed)
Note: Nucleases treatment might be needed to show if the TASQ staining induced by G4 ligand is G4- or rG4-specific. If nucleases treatment is required and CSK pre-extraction is possible, we recommend treating with nucleases during both pre-extraction and blocking steps (see General notes 1–4). Nevertheless, nucleases treatment is possible during the blocking step only.
1. If nuclease treatment is required during the pre-extraction step, prepare nucleases-supplemented CSK buffer (400 μL per well to be treated, as CSK treatment is performed twice) as follows (see General note 5):
a. For RNase-treated conditions, prepare CSK buffer supplemented with 0.3 mg/mL of RNase A.
b. For DNase-treated conditions, prepare CSK buffer supplemented with 1× TURBOTM DNase buffer and 10 U of TURBOTM DNase (final concentration 0.025 U/μL).
Caution: see General note 6.
2. If nuclease treatment is required during the blocking step, prepare the appropriate blocking buffer supplemented with nucleases (200 μL per well to be treated) as follows (see General note 5):
a. For RNase-treated conditions, prepare the appropriate blocking buffer supplemented with 0.3 mg/mL of RNase A.
b. For DNase-treated conditions, prepare the appropriate blocking buffer supplemented with 1× TURBOTM DNase buffer and 5 U of TURBOTM DNase (final concentration 0.025 U/μL).
Caution: We found that the appropriate blocking buffer (detailed in Materials) for MultiTASQ is BSA blocking buffer, and that for azMultiTASQ is FBS blocking buffer (see Troubleshooting 4, Figure 10).
Caution: See General note 6.
D. Post-fixation staining (Figure 3)
Figure 3. Representative images of MultiTASQ (A) and azMultiTASQ (B) clicked with AF594 (in red) as post-fixation G4 probes in HeLa cells treated or not for 4 h with 10 μM of pyridostatin (PDS), a well-characterized G4 ligand. Several conditions of fixative (MeOH or PFA) and pre-fixative (CSK supplemented or not with nucleases) techniques are represented to show versatility of TASQs staining (see General notes 1–4). Representative images are maximal z-projection of 10 deconvoluted z-stacks (total z = 3 μm). Similar brightness and contrast settings were applied for control and for PDS-treated images independently of fixative and pre-fixative conditions, allowing comparison between ctrl and PDS and optimization of signal pattern that can be observed. White dotted lines outline nuclei and scale bar (in light grey) stands for 20 μm in all images. Images were acquired with 60× oil-objective; experiments without TASQ (as exemplified in Figure 6) were performed as controls but not shown here.
1. After PDS treatment, remove medium (with aspiration system) and wash cells three times with 500 μL/well of cold DPBS (see General note 7).
Caution: If you plan to use pre-extraction (CSK) treatment with or without nuclease, follow steps D2–4, otherwise go to step D5.
Critical: All along the protocol, addition of buffers and compounds should be done with a low flow rate and on the edge of the well (especially before fixation) to avoid loss of cells.
Note: Removal of medium, buffers, and compounds should be done with an aspiration system to allow timeliness.
Note: We noticed that washing with cold DPBS (stored at 4 °C) before the fixation step allowed us to keep cells adherent.
Note: See General note 8.
See Troubleshooting 1.
2. Place the plate with the coverslips on ice. Remove DPBS and add 200 μL/well of ice-cold CSK (supplemented or not with nuclease) (see General note 2).
a. Incubate for 3 min on ice.
Critical: Inappropriate percentage of Triton X-100 in CSK buffer, time of incubation, and temperature of incubation could lead to loss of cells and should be optimized in a cell type–dependent manner (see Troubleshooting 2).
Note: In order to keep incubation times as accurate as possible, we recommend spacing the addition of CSK from 5–10 s apart between each well.
3. Gently remove the CSK (supplemented or not with nuclease) and repeat step D2.
Critical: Removal and addition of CSK could lead to cell detachment (see Troubleshooting 2).
4. Gently remove the CSK (supplemented or not with nuclease) and wash the cells once with 500 μL/well of cold DPBS.
Critical: After washing, fixation must be done as soon as possible.
5. If PFA-based fixation is used (see General note 2), prepare 4% PFA in DPBS (200 μL per well to be fixed) while cells are being washed in cold DPBS.
Critical: Solutions must be freshly prepared for each experiment from 16% PFA (see General notes 5 and 6).
Note: The percentage of PFA can be changed (typically 2%–4% PFA) if more suitable with previous experiments and/or immunodetection.
6. Gently remove the DPBS and fix the cells (see General note 2):
a. For 15 min at room temperature with 200 μL/well of 4% PFA solution, or
b. For 5 min on ice with 200 μL/well of 100% methanol.
Note: PFA fixation is mainly used after CSK pre-extraction.
Note: See General note 8.
7. Remove the fixative solution (with an aspiration system) and wash the cells three times with 500 μL/well of DPBS.
Pause point: After fixation and washing steps, you can keep the plate at 4 °C with 1 mL/well of DPBS for up to several days.
Note: See General note 8.
8. Remove the DPBS and permeabilize the cells (see General note 3):
a. For 10 min at room temperature with 200 μL/well of DPBS-Tx.
Note: If you have chosen the CSK pre-extraction protocol, permeabilization is not mandatory as Triton X-100 contained in CSK buffer already permeabilized cell membranes.
Note: See General note 8.
9. Remove the permeabilization solution and wash the cells three times with 500 μL/well of DPBS.
10. Remove DPBS and add 200 μL/well of the appropriate blocking buffer (supplemented or not with nuclease) (see General notes 5 and 6).
a. Incubate for 1 h at 37 °C.
Note: Preparation of blocking buffer supplemented with nuclease is detailed in section C.
11. Prepare a solution of 20 μM of TASQ in the appropriate blocking buffer (200 μL per well to be stained) (see General notes 5 and 6 and Recipes).
Caution: Supplementation with nuclease is not required at this step.
Note: Primary antibody can be diluted in this blocking buffer as immunodetection and TASQ incubation can be performed at the same time.
See Troubleshooting 4.
12. Remove the blocking buffer and add 200 μL/well of 20 μM of TASQ solution.
a. Incubate for 1 h at 37 °C.
Note: At this point, immunodetection is compatible; an incubation with primary antibody is possible in TASQ solution. However, we highly recommend testing if the blocking buffer allows for normal immunostaining.
13. Remove TASQ solution and wash five times with 500 μL/well of TBS-Tw.
Note: If TASQ staining is too low, reducing the number of washing steps is possible; however, background noise can be increased (see Troubleshooting 4).
14. Prepare the “click” solution (200 μL per well to be stained) (see General notes 5 and 6):
a. For MultiTASQ (for CuAAC) (see CuAAC mix Recipes).
b. For azMultiTASQ (for SPAAC) (see SPAAC mix Recipes).
Note: CuAAC “click” reaction with azMultiTASQ is also possible, but it requires alkyne-functionalized fluorophore; the selection of either TASQ depends on their availability given that MultiTASQ is readily accessible (being commercially available) while azMultiTASQ could be obtained upon request only.
15. Remove TBS-Tw and add 200 μL/well of “click” solution.
a. Incubate for 30 min at room temperature in the dark.
Caution: From this step, avoid exposure to light by keeping under an aluminum foil.
16. Remove the “click” solution and wash twice with 500 μL/well of TBS-Tw.
Note: After washing steps with TBS-Tw, incubation with a secondary antibody for immunodetection is possible.
Caution: After this step, follow section F for the end of the procedure.
E. Live TASQ staining (Figure 4)
Figure 4. Representative images of MultiTASQ (A) and azMultiTASQ (B) clicked with Alexa-Fluor 594 (in red) as live-cell treatment (1 h, 20 μM) after a 4 h treatment without (ctrl) or with 10 μM of pyridostatin (PDS), a well-characterized G4 ligand. Several conditions of fixative (MeOH or PFA) and pre-fixative (CSK supplemented or not with nucleases) techniques are represented to show the versatility of TASQs staining (see General notes 1–4). Representative images are maximal z-projection of 10 deconvoluted z-stacks (total z = 3 μm). Similar brightness and contrast settings were applied for control and PDS-treated images independently on fixation/pre-fixation conditions. The background was determined in extracellular spaces (three measurements), and background subtraction was applied to MeOH-fixed conditions (in A and B). White dotted lines outline nuclei, and scale bar (in light grey) stands for 20 μm for all images. Images were acquired with a 60× oil objective; experiments without TASQ (as exemplified in Figure 6 below) were performed as controls but are not shown here.
1. One hour before the end of the PDS treatment, treat each well with a final concentration of TASQ of 20 μM [i.e., add 5 μL of working solution of TASQ at 2 mM (see section A: Preparation of TASQs) directly in 500 μL/well DMEM (with or without PDS treatment)].
a. Incubate for 1 h at 37 °C.
Critical: Do not forget to keep TASQ-untreated wells for negative controls.
Note: Make sure the solution in the well is homogenous by gentle up-down pipetting.
Note: See General notes 7 and 8.
See Troubleshooting 1.
2. At the end of treatment, remove the medium and wash gently the cells three times with 500 μL/well of cold DPBS.
Caution: If you plan to use pre-extraction (CSK) treatment with or without nuclease, follow steps E3–5; otherwise, go to step E6.
Critical: All along the protocol, the addition of buffers and compounds should be done with a low flow rate and on the edge of the well (especially before fixation) to avoid loss of cells.
Note: Removal of medium, buffers, and compounds should be done with an aspiration system to allow timeliness.
Note: We noticed that washing with cold DPBS (stored at 4 °C) before the fixation step allowed us to keep cells adherent.
3. Place the plate with the coverslips on ice. Remove DPBS and add 200 μL/well of ice-cold CSK (supplemented or not with nuclease) (see General note 2).
a. Incubate for 3 min on ice.
Critical: Inappropriate percentage of Triton X-100 in CSK buffer, time of incubation, and temperature of incubation could lead to loss of cells and should be optimized in a cell type-dependent manner (see Troubleshooting 2).
Note: In order to keep incubation times as accurate as possible, we recommend spacing the addition of CSK from 5–10 s apart between each well.
4. Gently remove the CSK (supplemented or not with nuclease) and repeat step E3.
Critical: Removal and addition of CSK could lead to cell detachment (see Troubleshooting 2).
5. Gently remove the CSK (supplemented or not with nuclease) and wash the cells once with 500 μL/well of cold DPBS.
Critical: After washing, fixation must be done as soon as possible.
6. If PFA-based fixation is used (see General note 2), prepare 4% PFA in DPBS (200 μL per well to be fixed) while cells are being washed in cold DPBS.
Critical: Solutions must be freshly prepared for each experiment from 16% PFA (see General notes 5 and 6).
Note: The percentage of PFA can be changed (typically 2%–4% PFA) if more suitable with previous experiments and/or immunodetection.
7. Gently remove the DPBS and fix the cells:
a. For 15 min at room temperature with 200 μL/well of 4% PFA solution, or
b. For 5 min on ice with 200 μL/well of 100% methanol.
Note: PFA fixation is mainly used after CSK pre-extraction.
Note: See General note 8.
8. Remove the fixative solution (with aspiration system) and wash the cells three times with 500 μL/well of DPBS.
Pause point: After fixation and washing steps, you can keep the plate at 4 °C with 1 mL/well of DPBS for up to several days.
Note: See General note 8.
9. Remove the DPBS and permeabilize the cells (see General note 3):
a. For 10 min at room temperature with 200 μL/well of DPBS-Tx.
Note: If you have chosen the CSK pre-extraction protocol, permeabilization is not mandatory as Triton X-100 contained in CSK buffer already permeabilized cell membranes.
Note: See General note 8.
10. Remove DPBS and add 200 μL/well of the appropriate blocking buffer (supplemented or not with nuclease) (see General notes 5 and 6).
a. Incubate for 1 h at 37 °C.
Note: Preparation of blocking buffer supplemented with nuclease is detailed in section C.
11. Prepare the “click” solution (200 μL per well to be stained) (see General notes 5 and 6):
a. For MultiTASQ (for CuAAC) (see CuAAC mix Recipes).
b. For azMultiTASQ (for SPAAC) (see SPAAC mix Recipes).
Note: CuAAC “click” reaction with azMultiTASQ is also possible, but it requires an alkyne-functionalized fluorophore.
12. Remove TBS-Tw and add 200 μL/well of “click” solution.
a. Incubate for 30 min at room temperature in the dark.
Caution: From this step, avoid exposure to light by keeping under an aluminum foil.
13. Remove the “click” solution and wash twice with 500 μL/well of TBS-Tw.
Note: After washing steps with TBS-Tw, an incubation with a primary and secondary antibody for immunodetection is possible.
Caution: After this step, follow section F for the end of the procedure.
F. Nuclei counterstaining
1. Prepare the DAPI-staining solution (200 μL per well) with 1 μg/mL of DAPI in DPBS-Tx (see General notes 5 and 6).
2. Remove TBS-Tw and add 200 μL/well of the DAPI-staining solution.
a. Incubate for 15 min in the dark at room temperature.
3. Remove DAPI-staining solution and wash twice with 500 μL/well of TBS-Tw.
4. Keep coverslips in TBS-Tw in the dark until mounting.
Pause point: Stained coverslips can be kept in 1 mL of DPBS up to 24 h, in sealed wells, at 4 °C covered with aluminum foil before being mounted on a microscope glass slide.
G. Mounting coverslip on a microscope glass slide (Figure 5)
Note: Mounting coverslips can be tricky because of the risk of coverslip breakage, flip-over, air bubbles between the coverslip and the glass slide, etc. Here are a few tips.
Figure 5. Mounting of coverslips on a microscope glass slide. Cells were stained for 10 min with 0.057% of sulforhodamine B in order to give a better visualization of the mounting process (this should not be done to perform TASQs staining experiment). A. Vectashield mounting medium placed on microscope glass slide. B–E. Coverslip taken out of well. F–I. Coverslip placed on Vectashield with cells (F) facing upward, (G–H) facing toward, and (I) in mounting medium between the glass slide and coverslip. J. Excess of mounting medium wiped out. K. Coverslip sealed with nail polish.
1. Note the experiment ID (date, name, and place for each sample, fluorophores used, etc.) on the frosted part of the microscope glass slide.
2. Place a drop of Vectashield mounting medium (ca. 5–7 μL) for each coverslip to be mounted on the microscope glass slide.
Note: Tip needs to be cut due to Vectashield mounting medium viscosity.
Caution: Mounting medium not supplemented with DAPI provides better specific signal/noise ratio.
3. Take the coverslip out of the well (by pushing with a needle on one side of the well until the coverslip is vertical, or almost) and use a tweezer to put it on a paper towel with cells facing upward.
Note: To avoid breakage, do not push with the needle right in the middle.
4. Gently wipe the edge of the coverslip to remove the excess of buffer and put the coverslip with cells facing downward onto the Vectashield mounting medium.
Note: With a 13-mm diameter glass coverslip, up to four coverslips can be sealed on a classic microscope slide (26 × 76 × 1 mm).
Note: If the coverslip flips over and you do not know on which side of the coverslip the cells are, try to scrap with a needle (next to the edge of the coverslip) to see a scar indicating the side where cells are.
Note: To avoid air bubbles, we recommend placing one edge of the coverslip in the mounting medium and gently guiding it down rather than dropping it onto the glass slide.
5. Once all coverslips are mounted on the microscope glass slide, gently press on it with a paper towel to remove the excess of mounting medium and seal the edges of the coverslip with a small amount of transparent nail polish.
Pause point: Microscope glass slides can be kept at 4 °C in the dark before acquisition.
H. General guidelines for acquisition
Note: The type of microscope used for acquisition can also impact the quality of images. Even if the confocal laser scanning microscope (CLSM) is a golden standard, especially for colocalization studies, a wide-field microscope coupled with a deconvolution program can produce accurate images at a lower cost (e.g., we use Olympus IX83 with 60× oil objective).
Caution: The thickness of coverslips is also important for the quality of images; this feature is dependent on the microscope used (please refer to the manufacturer’s guide).
1. The focal point is easier to find with DAPI staining.
See Troubleshooting 3.
2. You have to set a negative control condition [e.g., without TASQ incubation but with fluorophore to be clicked (AF-az or DIBO-AF), Figure 6] to ensure the specificity of your staining and to set key features for acquisition (% excitation and exposure).
Note: If settings chosen for acquisition are too high, you will have nonspecific staining, thus enhancing background noise.
See Troubleshooting 4.
Figure 6. Determination of acquisition settings with negative control in post-fixation (A) or live-cell (B) TASQs staining. Staining with Alexa Fluor 594 (in red, without or with TASQs) was performed in HeLa cells treated or not with PDS and fixed with 4% PFA. Nuclei were counterstained with DAPI (in blue). Merged representative images are maximal z-projections of 10 raw z-stacks (total z = 3 μm). Similar brightness and contrast settings were applied to all images with the same fluorophore staining. Scale bar (in light grey) stands for 20 μm for all images. Images were acquired with a 60× oil objective.
3. A positive control must be chosen (e.g., with G4 ligand) where staining is maximal to set the optimal features of the acquisition (Figure 4B with azMultiTASQ).
Note: If these settings are too high, the signal will be saturated, which will not allow for proper quantification.
See Troubleshooting 4.
4. The same acquisition settings must be kept throughout all tested conditions to ensure a reliable comparison between the conditions.
5. For the same acquisition time, a choice must be made between quantity and quality: a high-throughput analysis (with a 20× air objective for example) with many cells to quantify does not allow for an accurate detection, while a high-resolution analysis (with a 60× oil objective for example) limits the number of images acquired (i.e., the number of cells to be analyzed).
6. Some of the technical limitations of the microscope can be bypassed using a deconvolution program that enhances the quality of your images (Figure 7); however, great care must be taken to ensure that this process does not create artifacts. We highly recommend referring to the supplier information sheet and reading this review for a better understanding of how convolution and deconvolution work [34].
Figure 7. Comparison of raw and deconvoluted images in post-fixation (A) or live-cell (B) TASQs staining. Staining with Alexa Fluor 594 (in red) was performed in HeLa cells treated or not with PDS, pre-extracted with CSK, and fixed with 4% PFA. Representative images are maximal z-projections of 10 z-stacks (total z = 3 μm). Similar brightness and contrast settings were applied for control and PDS-treated images. White dotted lines outline nuclei; scale bar (in light grey) stands for 20 μm. Images were acquired with a 60× oil objective; experiments without TASQ (as exemplified in Figure 6 above) were performed as controls but are not shown here.
Data analysis
We have written a user-friendly and open-access macro now available on GitHub (https://github.com/ICMUB/clickable-TASQ) that works with ImageJ [35] (or FIJI software, also in open-access) [36].
Note: Explanations of the different features used in the macro are available and described in the folder to be downloaded on GitHub.
Validation of protocol
This protocol was partially (unoptimized) used and validated in the following research article:
Rota Sperti et al. [27]. The multivalent G-quadruplex (G4)-ligands MultiTASQs allow for versatile click chemistry-based investigations. RSC Chem Biol 4: 456–465. DOI: 10.1039/d3cb00009e.
Another example of application, presented in Figure 8, shows how this protocol can be used to assess whether a candidate ligand modulates the G4 landscape in treated cells. We studied here the way PDS affects nuclear G4 foci in light of its established ability to stabilize DNA G4s (vide supra). To this end, HeLa cells were treated with 10 μM PDS for 4 h, and the quantification of the G4 foci was performed with MultiTASQ treatment either post-fixation (A–C) or upon live-cell incubation (D–F). The staining was performed by click chemistry (CuAAC) with AF-az after CSK pre-extraction and 4% PFA fixation. The images were then collected (A and D) and quantified using the developed macro (B, C, E, and F).
Figure 8. Quantification of MultiTASQ signal after treatment with 10 μM of PDS for 4 h or without (Ctrl), in either post-fixation (A–C) or live-cell treatment protocol (D–F); staining was performed after CSK pre-extraction and 4% PFA fixation. A and D. Representative field of quantified images (acquired with 60× oil objective and deconvoluted). B and E. Quantification of mean volume (per nucleus; in μm3) of MultiTASQ foci segmented with macro (large foci > 150 voxels; small foci ≤ 150 voxels; 150-voxels volume is determined to decipher nucleoli staining from other foci). Note: Different threshold values were used for segmentation as post-fixation and live-cell images have different intensities. C and F. Quantification of mean number (per nucleus) of MultiTASQ foci segmented with macro and with the same threshold values as for quantification of volume. B, C, E, and F. Mean values (lines and written on top of graph) and error bars (SEM) stand for one experiment with a number of nuclei analyzed n ≥ 144 (each nucleus is represented by a point). Note: We decided to quantify only nuclear staining as cytoplasmic staining was too faint.
These results were interpreted as follows: The post-fixation protocol showed that PDS was found to decrease the volume of nucleolar G4 foci (or large foci, B) without affecting their number (C) and to increase the volume of isolated G4 sites (or small foci, B) concomitantly decreasing their number (C). These mixed results were attributable to a rather elevated background fluorescence (grey dots in B, C), which originates in the widespread accumulation of MultiTASQ and/or of AF-az in fixed and permeabilized cells, and made it complicated to draw reliable conclusions. In contrast, the live-cell treatment protocol provided images in which the background was strongly decreased (E, F), which made the analysis of isolated G4 foci easier. The lack of large foci favors the detection of small foci: here, both the volume (ca. 4-fold) and number (ca. 40-fold) of the small foci increased upon PDS treatment, which is interpreted as a direct visualization of its ability to stabilize G4s and/or promote G4 folding in human cells.
General notes and troubleshooting
General notes
1. Some key features need to be optimized as they could impact TASQ staining:
a. This protocol is optimized for adherent cells; if you are using cells in suspension, technical adjustments must be done [37].
b. Type of cells: some cells are more fragile than others and need optimization for pre-fixation treatments and/or fixation protocols (see Troubleshooting 1 and 2).
2. Fixation
a. Pre-fixation techniques: this technique, as well as fixation techniques, can either increase the quality of visualization or induce artifacts.
• CSK pre-extraction improves staining by removal of soluble proteins [38]; only proteins attached to macro-molecular structures (such as cytoskeleton, chromatin, etc.) are maintained, which decreases the background noise. However, this technique has some limitations (see Troubleshooting 2).
• Nuclease (DNase and/or RNase) supplemented pre-fixation treatment, whenever possible, improves the efficiency of DNA and/or RNA digestion compared to post-fixation treatment.
b. Fixation techniques: a poor fixation can lead to loss of material/signal while an over-fixation can lead to artifacts [39].
• Paraformaldehyde (PFA) is a crosslinking fixative reagent commonly used at 4%; however, nuclear proteins are more efficiently fixed with 2% and a longer fixation time. The formation of intermolecular bridges by PFA allows for the fixation of soluble proteins but can also mask some antigenic sites.
• Methanol is a dehydrating fixative reagent that should be used at low temperature to avoid artifacts due to denaturation and/or precipitation of biomolecules. This technique is not suitable for fixation of soluble or state-modified (such as phosphorylation) targets.
3. Permeabilization techniques: methanol and various detergents (such as Triton X-100 and Tween 20) are used to create different sizes of pores in the plasma membrane, but it can also lead to loss of membrane proteins and organelles targets for live-cell staining experiments.
4. The selection of TASQs and related fluorophores:
a. TASQ staining: versatile “clickable” TASQ can be used as a live-cell G4 probe for the detection of G4s in a cellular context. During live-cell treatment, the concentration of TASQ, duration of treatment, and competition with other G4 ligands (such as PDS) with a better affinity than TASQ (APPKD in the μM range, determined by fluorescence quenching assay) [27,40] may displace TASQs if the order of co-treatment is not properly conducted; it should be optimized to avoid misinterpretations.
b. Using TASQ as a post-fixation probe allows for more intense G4 staining, which is not related to TASQ availability issues but to the fact that post-fixation staining allows for the detection of fixed native and PDS-induced G4s. These differences between post-fixation and live-cell staining could explain the differences between the images seen in Figures 3 and 4. Additionally, a short-time live-cell TASQ incubation can lead to high extracellular background noise, which can be computationally removed after image acquisition.
c. We recommend the use of Alexa-Fluor (AF) fluorescent compounds (for “click” reactions and, if needed, for immunodetection with secondary antibody) as bleaching is reduced with these fluorophores, thus increasing the quality of their detection. However, the use of AF594 is not mandatory and could be adapted, particularly when immunodetection is performed given that the same results were obtained with other AF fluorophores (including AF488 and AF647).
d. Co-immunostaining: if co-immunostaining is required, be sure fluorophore multiplexing is possible and the fixation protocol is compatible with antibody recognition of targeted antigen. Fluorescence overlapping is still possible (resulting in noise); therefore, appropriate controls (i.e., the fluorophore to be clicked without TASQ incubation and the secondary antibody without primary antibody) must be performed to ascertain the specificity of the observed signals.
5. We recommend preparing a mix with an extra volume (typical example of mix: required volume per well × number of well to be treated + 100 μL).
6. We highly recommend preparing fresh dilutions for some compounds [e.g., buffers supplemented with nucleases (if needed), PFA solutions, TASQs in recommended buffers (20 μM), buffers for click chemistry].
7. Agitation during any washing steps is not recommended as it can lead to loss of cells (especially before fixation), and it does not improve signal/noise ratio.
8. We highly recommend following the morphology of cells (Figure 9) after critical steps of pre-extraction, fixation, and permeabilization with a brightfield or phase-contrast microscope. These steps are not sensitive to light exposure and therefore can be done without any impact on TASQ-mediated G4 detection.
a. During fixation, you can check on the morphology of cells after CSK pre-extraction with a brightfield or phase-contrast microscope. It is important to note that this can be done only during fixation to avoid cell detachment with plate movements. Cells look smaller and cell membranes have (almost) disappeared, and only nucleus membranes are visible.
b. If you used only PFA (without pre-extraction), you will be able to see a difference before and after permeabilization on the morphology of fixed cells, particularly on cell membranes.
Figure 9. Following morphology (with brightfield microscopy) of HeLa cells before Alexa Fluor staining. A. In DMEM, before the first PBS washing step. B. After the first PBS washing step, before fixation. C–G. Without pre-extraction; H–L. after pre-extraction with CSK buffer. C and H. Two minutes after the beginning of PFA 4% fixation. D and I. After fixation and washing with PBS. E and J. Two minutes after the beginning of permeabilization. F and K. After permeabilization and washing with PBS. G and L. In mounted coverslips. Representative images are one z-stack acquired with a 10× air objective. Scale bar (in black) stands for 100 μm.
Troubleshooting
Problem 1: Low number of cells at the end of treatment (i.e., before pre-extraction or fixation).
Possible cause: Not enough cells seeded.
Solution: Try several dilutions; the number of cells can be based on confluency in a T75 flask, i.e., confluency, number of cells, doubling time, and the surface of seeding can be used to have an idea of the number of cells to be seeded.
For example, if you have 12 × 106 cells with 70% confluency in a T75 flask (surface of flask: 75 cm2), you will have 70% confluency with 300,000 cells per well in a 24-well plate (surface per well: 1.9 cm2). To seed cells 24 h before treatment (with short-time treatment ≤ 4 h) with a doubling time of 16 h and obtain 70% confluency on the day of treatment, you will need:
300000
2
24
/
16
≈
106000
cells per well.
Possible cause: Cells are not adherent to glass coverslips.
Solution: Try to seed cells for a longer time before treatment; we tried for 40 h before treatment with MCF7 (cell type used only in Figure 10) to allow them to recover adhesion and normal phenotype.
Solution: Try with coated coverslips to increase cell adhesion.
Possible cause: Treatment is too cytotoxic and induces cell death and/or limits cellular adhesion.
Solution: Try with a lower dose or duration of treatment to reduce cytotoxicity; if you cannot change these, try with coated coverslips.
Problem 2: Loss of cells after pre-extraction or fixation.
Possible cause: Washing steps before fixation were too harsh.
Solution: Aspirate and dispense washing buffer on the edge of the wells to avoid high flow pressure. As liquid velocity (therefore fluid pressure) is inversely proportional to the diameter, use bigger tips (for instance, to dispense 200 μL, it is better to use a P1000 than a P200) or cut the end of tips.
Solution: Dispensing the washing buffer on the edge of wells also avoids drops falling in the middle of the well, where the most interesting area to look at is likely to be.
Possible cause: Pre-extraction is too harmful for cells.
Solution: As CSK buffer is highly hypotonic and detergent, we highly recommend being accurate with incubation times with this buffer; even 30 s too long on each incubation can be detrimental.
Solution: If there is still a loss of cells during this step, then the incubation time and the number of incubations with CSK buffer can be decreased to avoid cells’ detachment.
Solution: The percentage of Triton X-100 must be optimized depending on the cell line: a reduction of up to 0.3% of Triton X-100 does not affect the quality of the pre-extraction (observed with MCF7 and MIA PaCa-2 cell lines, data not shown).
Solution: If all these solutions do not work, it might happen that cells need more time to recover an unstressed state and normal phenotype; try to seed cells for a longer time before treatment.
Note: Seeding more cells is not a good solution as entire patches of cells can be detached if adhesion between cells is stronger than the adhesion of cells to the glass coverslip.
Possible cause: Methanol fixation has decreased the number of cells.
Solution: Methanol has to be cold: keep it at -20 °C. If methanol is the only fixative technique used, you can place the plate at -20 °C during fixation (which is not compatible with PFA fixation).
Solution: Shortened time of fixation with methanol.
Problem 3: DAPI staining is blurry, or nuclei are overlapping
Possible cause: Too many cells seeded (confluency on microscope field ≥ 80 %).
Solution: Decrease the number of cells.
Possible cause: Several layers of cells or aggregates of cells (see Figure 10).
Solution: Before seeding, use trypsin for a longer time (but without any effect on cell viability) and make sure cells are perfectly resuspended. If not, it can be seen during counting.
Solution: For seeding, prepare an intermediate dilution to allow the dislocation of cell aggregates.
Solution: Vortex solution containing cells to be seeded for 5–10 s (if vortex can be used with your cell line; using vortex is compatible with HeLa cells) and directly seed cells.
Solution: Be gentle with the homogenization of cells in your plate to avoid having all cells localized at one edge of the well.
Possible cause: Cells above coverslips (trapped between the coverslip and plastic plate during incubation at 37 °C and fixed on coverslip).
Possible cause: Condensation when returning to room temperature after storage at 4 °C.
Solution: Gently wipe the top of coverslips with Kimwipes without disturbing the sealing with nail polish (once mounted and sealed on microscope glass slide).
Problem 4: Signal/noise ratio enhancement
Possible cause: Nonspecific localization of clickable Alexa Fluor.
Solution: Increase the number of washing steps and/or wash with a more stringent buffer (increasing detergent percentage).
Solution: Decrease the concentration of Alexa Fluor used for click reaction.
Solution: Change the blocking buffer (see Figure 10).
Possible cause: TASQ staining has faded from one experiment to another (from reproducible experiments or from post-fixation to live cell experiments)
Solution: Prepare a new 2 mM dilution of TASQ from the stock solution of TASQ (10 mM) to ensure TASQ integrity.
Solution: For live staining, the time of TASQ incubation can be increased in order to maximize the entry of compounds.
Note: We highly recommend setting all technical considerations on post-fixation experiments as staining is easier to observe in this condition.
Figure 10. Optimization of background noise induced by the use of DIBO-AF594 for azMultiTASQ staining in post-fixation. Staining with Alexa Fluor 594 (in red) was performed in MCF7 cells fixed with PFA 4%. Representative raw images were acquired with a 20× objective with the same acquisition parameters for all conditions (independently in A and B, from two independent experiments). The same brightness and contrast (B/C) settings (either low or high B/C) were applied for all conditions. White lines outline nuclei, and scale bar (in light grey) stands for 20 μm. A. Comparison of click reaction in order to decrease nonspecific cellular AF594 staining: with AF-az (for MultiTASQ) and with DIBO-AF (for azMultiTASQ) performed in the same conditions (with BSA-based blocking buffer only during first incubation ± TASQs and PBS as click reaction buffer), with DIBO-AF click reaction performed in BSA-based blocking buffer and with DIBO-AF click reaction performed in 1% FBS-based blocking buffer (with 1% FBS-based blocking buffer for first incubation ± TASQs). B. Comparison of click reaction in order to decrease nonspecific extracellular AF594 staining with 1% or 2% FBS-based blocking buffer, consistent with the protocol we recommended.
Acknowledgments
This work was supported by the CNRS and the Agence Nationale de la Recherche (ANR-22-CE44-0039-01, InJUNCTION). The authors are grateful to Francesco Rota Sperti, Sandy Raevens, and Dr. Ibai E. Valverde (ICMUB, Dijon, France) for the preparation of TASQ tools. This protocol was adapted and modified from Rota Sperti et al. [27], DOI: 10.1039/d3cb00009e.
Competing interests
The CNRS has licensed MultiTASQ to Merck KGaA for commercialization.
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Article Information
Publication history
Received: Oct 25, 2024
Accepted: Dec 18, 2024
Available online: Jan 19, 2025
Published: Feb 5, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Molecular Biology > RNA > RNA labeling
Cell Biology > Cell imaging > Live-cell imaging
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5,210 | https://bio-protocol.org/en/bpdetail?id=5210&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
A Bio-protocol resource
Peer-reviewed
Real-time IncuCyte® Assay for the Dynamic Assessment of Live and Dead Cells in 2D Cultures
AG Arlene K. Gidda
SC Suganthi Chittaranjan
SG Sharon M. Gorski
Published: Vol 15, Iss 3, Feb 5, 2025
DOI: 10.21769/BioProtoc.5210 Views: 77
Reviewed by: Ralph Thomas BoettcherShun Yu Jasemine Yang Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Journal of Cell Science Oct 2023
Abstract
Cell viability and cytotoxicity assays are commonly used to investigate protein function and to evaluate drug efficacy in cancer and other disease models. Cytotoxicity is the measure of dead or damaged cells and is often quantified using assays based on cellular characteristics such as membrane integrity or mitochondrial metabolism. However, these assays are typically limited to endpoint analysis and lack emulation of physiological conditions. The IncuCyte Live and Dead Cell assay described here leverages common cell permeability methodologies but uses fluorescence microscopy channels to image both live and dead cells over time and phase microscopy channels to measure confluency. Cytotox green reagent is a cell membrane–impermeable dye that can only be taken up by cells with poor cell membrane integrity. NucLight rapid red dye is a cell membrane–permeable nuclear dye that can be taken up by all cells. Based on dye uptake and fluorescence intensity, the IncuCyte software can be used to analyze images for live and dead cell detection and quantification. Phase microscopy is used to determine confluency and can be further quantified using the IncuCyte software. We provide an application of this assay, using it to calculate IC50 and EC50 values for the assessment of drug efficacy.
Key features
• Quantify live and dead cells over time.
• Determine drug IC50 and/or EC50 in 2D cell cultures.
• This protocol requires the instrument IncuCyte® S3 (or SX5) Live-Cell Analysis system and corresponding software.
Keywords: IncuCyte Cytotoxicity assay 2D cell culture Chloroquine MIA Paca-2 Cytotox Green NucLight Rapid Red
Graphical overview
Background
Under normal physiological conditions, many cell types undergo turnover to help maintain a balance between cell death and cell survival. However, in cancer contexts, there is an imbalance that favors cell survival or proliferation over cell death. Cancer cell death is an important area of cancer research, especially in drug discovery [1]. High-throughput, sensitive, and robust assays are necessary for screening, testing, and comparing the effectiveness of candidate drugs [1]. The effectiveness of drugs is commonly measured by the half-maximal inhibitory concentration (IC50) or the half-maximal effective concentration (EC50) [2]. The IC50 is the concentration at which half of the measured response is inhibited, while the EC50 is the concentration at which half of the maximum response is induced [2]. One way of measuring drug effectiveness in cancer models is by determining how well a drug induces cytotoxicity [1]. Cytotoxicity can be evaluated via various indirect measures of cell death including cell permeability (trypan blue dye exclusion assay), metabolism (tetrazolium reduction assay), protease activity, ATP detection, and many more [3,4]. In addition to these assays being indirect measures of cell death, other drawbacks typically include the lack of physiological conditions and restriction to end-point analyses [5]. One way to overcome these limitations is by using the IncuCyte Live and Dead Cell assay.
The IncuCyte Live and Dead Cell assay employs common cell permeability techniques, but in a capacity that allows for real-time and live-cell quantification. Cytotox green reagent is a membrane-impermeable dye that is only able to enter cells with poor membrane integrity, hence serving as a cell death marker. NucLight rapid red dye is a membrane-permeable DNA dye that labels all cell nuclei. Residing in a tissue culture incubator, the IncuCyte® Live-Cell Analysis system can capture images at fixed time intervals, enabling the quantification of dead vs. live cells using the IncuCyte software. Live and dead cell quantification is used to calculate the IC50 and/or EC50. Changes in cell morphology, cell migration, and cell–cell clustering interactions throughout the run can also be quantified.
In this protocol, we describe stepwise how to perform the IncuCyte Live and Dead Cell assay, providing an example of how it can be used to determine the IC50 and EC50 of chloroquine (CQ), a late-stage autophagy inhibitor, in the pancreatic ductal adenocarcinoma cell line, MIA PaCa-2. A modified version of this protocol was used in Sathiyaseelan et al. [6].
Materials and reagents
Biological materials
1. MIA Paca-2 cells (ATCC, catalog number: CRL-1740)
Reagents
1. Dulbecco’s modified Eagle medium (DMEM) (Thermo Fisher Scientific, Gibco, catalog number: 11995-065)
2. Fetal bovine serum (FBS) (Thermo Fisher Scientific, Gibco, catalog number: 12483-020)
3. HEPES (Thermo Fisher Scientific, Gibco, catalog number: 15630-080)
4. Insulin solution from bovine pancreas (Sigma-Aldrich, catalog number: 10516)
5. MEM non-essential amino acids (NEAA) (Thermo Fisher Scientific, Gibco, catalog number: 11140-050)
6. 0.25% Trypsin-EDTA phenol red (Thermo Fisher Scientific, Gibco, catalog number: 25200056)
7. PBS, pH 7.4 (Thermo Fisher Scientific, Gibco, catalog number: 10010-049)
8. EVETM slides and 0.4% trypan blue solution (NanoEnTek, catalog number: EVS-050)
9. Chloroquine diphosphate (Sigma-Aldrich, catalog number: C6628-25G)
10. Staurosporine (Sigma-Aldrich, catalog number: S6942)
11. IncuCyteTM NucLight rapid red dye for live-cell nuclear labeling (Sartorius, catalog number: 4717)
12. IncuCyteTM Cytotox green reagent for counting dead cells (Sartorius, catalog number: 4633)
Solutions
1. Full media (see Recipes)
2. Chloroquine (CQ) stock solution (see Recipes)
Recipes
1. Full media (500 mL)
Note: Store the media for up to 2–3 weeks at 4 °C. Warm to 37 °C before using.
Reagent Stock concentration Final concentration Quantity or Volume
DMEM n/a n/a 439.75 mL
FBS 100% 10% 50 mL
NEAA 100× 1× 5 mL
HEPES 1 M 10 mM 5 mL
Insulin 10 mg/mL 5 μg/mL 250 μL
2. Chloroquine (CQ) stock solution (1.9 mL)
Note: The stock is 100 mM and is stable for at least 1 year at -20 °C. For experiments, dilute to 2 mM (5 µL of 100 mM stock + 245 µL media) in full media and make further dilutions accordingly in full media.
Reagent Final concentration Quantity or Volume
Chloroquine diphosphate (MW: 515.86) 100 mM 0.1 g
1× PBS n/a 1.9 mL
Laboratory supplies
1. T75 or T25 flasks (VWR, catalog number: BD353136 or 29185-300)
2. 70% ethanol (HealthCare Plus, catalog number: NPN 00825751)
3. 96-well tissue culture plates (Corning, Falcon, catalog number: 353072)
4. Sterile 15 mL tubes (Corning, Falcon, catalog number: 352096)
5. 10 mL pipette (Corning, catalog number: 4488) and pipet-aid (or aspirator)
6. 5.0 mL screw cap MacroTubes (MTC Bio Incorporated, catalog number: C2540)
7. Syringes for DISTRIMAN Mini ST 1,250 μL (Gilson, catalog number: F164140)
Equipment
1. DISTRIMAN repetitive pipette (Gilson, model: F164001)
2. EVE automatic cell counter (NanoEnTek, catalog number: EVE-MC)
3. Autoflow IR Direct Heat CO2 incubator (NuAire, model: NU-5510E)
4. IncuCyte S3 live-cell analysis system (Sartorius, IncuCyte S3)
5. Tissue culture hood (Microzone Corporation, model: BK-2-4)
6. Vortex Genie 2 (Scientific Industries, model: SI-0236)
Software and datasets
1. IncuCyte software 2022B (IncuCyte, 2022)
2. Excel 2016 (Microsoft, 09/22/2015); requires license
3. Prism v10.2.3 (GraphPad, 04/21/2024); requires license
4. BioRender (https://www.biorender.com/). The following figures were created using BioRender: Graphical overview: BioRender.com/c77h076
Procedure
A. Cell culturing
1. Grow MIA Paca-2 cells in full media in T75 (or T25) flasks in a humidified incubator at 37 °C with 5% CO2 until they reach 80%–90% confluency. For the MIA Paca-2 cell line, we typically split cells at a ratio of 1:3 when they reach 80%–90% confluency, which takes 3–4 days.
2. Sterilize the tissue culture hood with 70% ethanol. Spray down the container with tubes, pipettors, tips, and media bottle with 70% ethanol and place them in the hood for 20 min. Bring MIA Paca-2 cells inside the hood.
3. Remove media with an aspirator or a 10 mL pipette and wash cells three times with 6 mL of 1× PBS. Add 1 mL of 0.25% trypsin-EDTA per T75 flask (500 μL per T25). Incubate flasks at 37 °C in the incubator for 2–3 min until the cells completely detach.
4. Tap the flask to dislodge cells completely and prepare cell suspension in the tissue culture hood. Add 6–8 mL full media as required and pipette cells 5–10× with a 10 mL pipettor to resuspend cells. Transfer cell suspension to a 15 mL Falcon tube.
5. Gently vortex cells at a low speed and take cell counts with a hemocytometer or a cell counter using a cell viability dye like trypan blue as per the manufacturer’s instructions.
Note: Vortexing may not apply to all cell lines.
6. In a 15 mL Falcon tube (or 50 mL as required), make 10–15 mL of 5 × 104 cells/mL (50 cells/μL) suspension in the full media. Plate 100 μL of 50 cells/μL with a repeater pipettor in each well of a 96-well plate.
Note 1: We pipette 2× 50 µL per well (not 1× 100 µL), which gives a more equal distribution of cells per well.
Note 2: Cell numbers should be empirically determined for each cell line based on proliferation rates and the total duration of the experiment. Initial well confluency of 30% is recommended by Sartorius for cytotoxicity assays. Avoid high seeding densities because contact inhibition slows cell growth and makes it difficult to distinguish cells using imaging analysis.
7. Dilute NucLight rapid red dye (1:500 final well dilution) and Cytotox green reagent (250 nM final well concentration) together in full media and dispense 50 μL per well (at this point, the total well volume should be 150 μL). The final dilutions or concentrations should be calculated based on the final well volume of 200 μL.
Note 1: The concentrations of NucLight red and Cytotox green dyes, added together in a volume of 50 µL, should be calculated based on a final well volume of 200 µL. The same applies for the drug (step A8).
Note 2: Dyes can also be added after the cells adhere, but care should be taken not to disturb the cells. The dyes are nontoxic and incorporate readily into DNA; they typically remain detectable for 72 h or more, but dye retention is cell line dependent.
Note 3: According to Sartorius, cells are labeled immediately upon the addition of NucLight red.
8. Incubate plates at 37 °C in the incubator for 2–3 h until cells adhere. Meanwhile, prepare the drug for treatment (e.g., chloroquine) at required concentrations. Refer to Recipe 2 for preparing chloroquine stocks and further dilutions in full media. Add 50 μL of drug volume to the respective wells taking care not to disturb the cells (at this point, the total well volume should be 200 μL). Place the 96-well plate lid back on the plate and gently tap the plate to mix the reagents well.
B. Setting up IncuCyte protocol
1. Place the 96-well plate into the IncuCyte S3 instrument located in a humidified incubator at 37 °C with 5% CO2.
2. Let the plate sit in the IncuCyte for at least 30 min before starting the program to allow adjustment to the incubator temperature. Scans can be taken at any desired time interval. We generally record 4–5 images (recording 3 images also works well and saves scan time) per well every 4 h (or 2 h as required) at 10× objective to monitor cell proliferation and cell death for up to 72 h. Include negative (e.g., untreated or vehicle) and positive (e.g., staurosporine for cell death [7]) controls. Full media was the vehicle control used in our experiments. In 96-well plates, we exclude the outer wells from the experiment and fill them with media to avoid evaporation effects. We typically have three technical replicates per treatment.
3. Follow the steps as indicated in Figure 1 to start the run for IncuCyte S3 (or follow instructions for the other models or instruments as per the manufacturer’s instructions). We use default settings for acquisition times for 10× objective scans. We prefer to create a plate map for our runs (Figure 2) using the IncuCyte plate map editor software, but it is not necessary.
Figure 1. Key steps to start a run. A. Click on the “plus” sign to schedule your run. B. Select scan settings including image channels and objective. C. Select wells to scan and the number of images per well. D. Name run and provide plate map (this can be added after the run is complete). E. Define an analysis and spectral unmixing (this can also be done after the run ends).
Figure 2. Example IncuCyte plate map. The map depicts well location of MIA Paca-2 cells untreated (red) or treated with chloroquine (CQ; yellow; increasing shade corresponds to increasing drug concentration) or staurosporine (green). Three replicate wells were used for each condition. Creating a plate map facilitates data analysis as the software recognizes the identity of each well and labels the data correspondingly. Compounds, cells, and growth conditions can be added to the plate map description on the left.
Data analysis
A. Creating an analysis definition
1. For data analysis, we use the integrated algorithms of the IncuCyte software S3 (for details, refer to the User Manual available in the top left corner under the Help tab) to calculate NucLight red–positive (red fluorescence, total cells) and Cytotox green–positive (green fluorescence, dead cells) cells. Representative IncuCyte images for each treatment condition after 72 h are shown in Figure 3.
Figure 3. IncuCyte images. IncuCyte images showing NucLight red–positive and Cytotox green–positive cells taken after 72 h of run time at 10× objective, treated with 0.0005 mg/mL of staurosporine and varying concentrations of chloroquine (CQ). Staurosporine is the positive control, and full media is the negative control.
2. We use a basic analyzer for this assay and take images with phase (optional), red and green channels, and include overlap (green and red channel). Spectral unmixing was set at 5%–8% red removed from green to better represent the distribution of both green and red channels. Sartorius recommends 2%–8% red removed from green. The value is incrementally adjusted until the red fluorophore signal is not detected in the green channel. An example analysis window can be seen in Figure 4A. For MIA PaCa-2 cells, the red, green, and overlap counts were set as follows:
Red object count (total nuclei) parameters: Top-hat segmentation with 100 μm radius and 1 threshold; edge split on with edge sensitivity set to -30 (Figure 4B).
Green object count parameters: Top-hat segmentation with 100 μm radius and 2 threshold; edge split on and edge sensitivity set to -30. Adjust size set to -3. Dead cells were defined as double-positive red+/green+ objects and were identified by their overlap (overlap area min 50 mm2) (Figure 4C). These settings were optimized on a representative set of images.
Note: The settings should be adjusted for each adherent cell type. The User Manual describes the setting parameters in detail. Once settings for a particular cell line have been defined, the same settings should be used in all subsequent experiments for that cell line.
Figure 4. Defining an analysis. A. An example of an IncuCyte analysis window. Settings can be adjusted and previewed to determine the appropriate capture of confluency, red, green, and overlap masks. B. Mask for red object count (total cells). C. Overlap mask of red and green object counts (dead cells).
3. The red fluorescence, green fluorescence, and overlap counts are exported for further analysis.
B. Live and dead cell time-lapse analysis using Microsoft Excel
1. Using Excel, determine relative total cell (red cells relative to the 0 h time point, the time of treatment initiation), relative confluency (relative to the 0 h time point), and relative dead cell [yellow (green+red) cells relative to total cells] kinetics for up to 72 h. Live cell kinetics can be determined by subtracting dead cells (yellow cells) from total cells (red cells). Refer to Table 1 for further details.
Note: Relative cell death is more informative than just dead cell number since total cells also change over time.
Table 1. Description and formulas for calculating relative total cells, relative dead cells, relative live cells, and relative confluency
Parameter Description/Formula
Relative total cells Red cells per mm2 normalized to 0 h time point
Relative dead cells Yellow (red and green overlay count) cells per mm2 divided by red cells per mm2
Relative live cells Red cells per mm2 minus yellow cells per mm2 normalized to 0 h time point
Relative confluency Confluency (area of vessel covered in cells) normalized to 0 h time point
*Note: If presenting data as percentages, then values should be multiplied by 100.
2. Using GraphPad Prism software, graph relative total cells for each treatment concentration (Figure 5A). Repeat this for dead cells (Figure 5B), live cells (Figure 5C), and confluency (Figure 5D).
Figure 5. Graphing data and determining IC50/EC50. MIA Paca-2 cells were treated with increasing concentrations of chloroquine (CQ) for 72 h. Errors bars represent SEM. A. Total cell count normalized to the first time point. B. Dead cells relative to total cell fraction (dead/total cells). C. Live (total-dead) cell count normalized to the first time point. D. Phase confluency (determined by phase contrast microscopy channel images) normalized to the first time point. E. IC50 and EC50 of chloroquine based on total cell–normalized AUC and dead cell AUC. For the IC50, curve [Inhibitor] vs. normalized response was used for curve fitting on GraphPad. For the EC50 curve, [Agonist] vs. response (three parameters) was used for curve fitting. F. IC50 and EC50 based on confluency-normalized AUC and dead cell–AUC, respectively. For IC50, [Inhibitor] vs. normalized response–variable slope was used.
C. GraphPad Prism analysis
1. Next, using total and dead cell kinetics, determine IC50 and EC50 values. In GraphPad Prism (v10.2.3), under Analysis, click on Analyze and choose Area under curve (AUC).
2. Export treatment concentrations and AUCs for each treatment from both total and dead cell analyses to a new XY table, respectively. Just as before, under Analysis, click on Analyze, but this time choose Nonlinear regression. To determine IC50 values, use total cell data: choose dose-response-inhibition followed by [inhibitor] vs. normalized response to “curve-fit” (Figure 5E). (Note: We used normalized AUC and did not use logarithmic values of treatment concentrations because our drug concentration range was relatively small.) Curve-fitting should be chosen according to the data used and depends on the slope of the curve and whether the data is normalized and/or logarithmic. Please see Help in GraphPad software for details. This analysis was also repeated with confluency AUC data and [Inhibitor] vs. normalized response, in which the variable slope was used to curve fit (Figure 5F).
Note: The IC50 values determined by relative total cell and relative confluency are comparable.
3. Likewise, use dead cell AUCs for each treatment and determine EC50 values: use dose-response stimulation followed by [Agonist] vs. response (three parameters) to curve-fit (Figure 5E, F). The choice for curve-fit depends on the response data and the type of data (i.e., logarithmic, normalized, etc.) you use, as indicated above.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article:
• Sathiyaseelan et al. [6]. Loss of ATG4B and ATG4A results in two-stage cell cycle defects in pancreatic ductal adenocarcinoma cells. Journal of Cell Science (Figure 8, panel H).]
General notes and troubleshooting
General notes
This 2-color live and dead cell assay provides a multiplex way to analyze the viability and cytotoxicity effects of various treatments from the same well. Live and dead cell data from the same wells can be analyzed to compare IC50 and EC50 values (Figure 5E, F), providing insight into both the viability and cytotoxicity effects of the drug treatment. An advantage of the 2-color live and dead cell assay compared to the confluency assay is that it directly quantitates all cells and distinguishes between those live and dead. The confluency assay includes dead cells that are still attached to the plate (but does not identify them as dead) and thus can overestimate viability in such instances. Similar live and dead cell assays can be performed using alternate dead cell dyes available. For example, instead of Cytotox green dye, a caspase-3/7 green dye can be used to measure apoptotic cells.
Troubleshooting
Problem 1: Some cells do not take up NucLight red or efflux NucLight red within the experimental timeframe.
Solution(s): In these cases, confluency can be used to estimate total cells and determine IC50 values. Alternatively, cells stably expressing nuclear markers [histone B2 (H2B)-mCherry, IncuCyte® NucLight lentivirus system] can be used for total or live cell counts.
Problem 2: Air bubbles in the sample wells can cause images to be distorted.
Solution(s): Use a squeeze bottle with 70% ethanol with the straw removed to pass air above the wells and pop the air bubbles.
Problem 3: Dyes can clump together and create background signals that interfere with analysis.
Solution(s): Make sure to resuspend dyes thoroughly before adding them to wells.
Acknowledgments
We thank J. Chan and G. Samarasekera for valuable comments and feedback. This work was supported by the Canadian Institutes of Health Research (PJT-159536) and the Canadian Cancer Society (grant #707495). This protocol was adapted from the methods section of the study by Sathiyaseelan et al. [6].
Competing interests
The authors declare they have no conflict of interest.
References
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Caldwell, G. W., Yan, Z., Lang, W. and A. Masucci, J. (2012). The IC50 Concept Revisited. Curr Top Med Chem. 12(11): 1282–1290.
Adan, A., Kiraz, Y. and Baran, Y. (2016). Cell Proliferation and Cytotoxicity Assays. Current Pharm Biotechnol. 17(14): 1213–1221.
Khalef, L., Lydia, R., Filicia, K. and Moussa, B. (2024). Cell viability and cytotoxicity assays: Biochemical elements and cellular compartments. Cell Biochem Funct. 42(3): e4007.
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Sathiyaseelan, P., Chittaranjan, S., Kalloger, S. E., Chan, J., Go, N. E., Jardon, M. A., Ho, C. J., Hui, T., Xu, J., Chow, C., et al. (2023). Loss of ATG4B and ATG4A results in two-stage cell cycle defects in pancreatic ductal adenocarcinoma cells. J Cell Sci. 136(19): e260644.
Su, X., Wang, J., Jiang, L., Chen, Y., Lu, T., Mendonca, M. S. and Huang, X. (2021). PCNA inhibition enhances the cytotoxicity of β-lapachone in NQO1-Positive cancer cells by augmentation of oxidative stress-induced DNA damage. Cancer Lett. 519: 304–314.
Article Information
Publication history
Received: Aug 30, 2024
Accepted: Dec 30, 2024
Available online: Jan 19, 2025
Published: Feb 5, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
How to cite
Category
Cancer Biology > Cell death > Cell biology assays
Cell Biology > Cell viability > Cell death
Cell Biology > Cell-based analysis
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5,211 | https://bio-protocol.org/en/bpdetail?id=5211&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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This is an In Press version of the protocol that has not yet been assigned to an issue.
Peer-reviewed
Nuclei Isolation From Murine and Human Periosteum For Transcriptomic Analyses
SP Simon Perrin
CG Cassandre Goachet
ME Maria Ethel
YH Yasmine Hachemi
CC Céline Colnot
In Press, Available online: Jan 22, 2025
DOI: 10.21769/BioProtoc.5211 Views: 22
Reviewed by: Samantha HallerVishal Nehru Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in eLIFE Dec 2024
Abstract
Bone repair is a complex regenerative process relying on skeletal stem/progenitor cells (SSPCs) recruited predominantly from the periosteum. Activation and differentiation of periosteal SSPCs occur in a heterogeneous environment, raising the need for single cell/nucleus transcriptomics to decipher the response of the periosteum to injury. Enzymatic cell dissociation can induce a stress response affecting the transcriptome and lead to overrepresentation of certain cell types (i.e., immune and endothelial cells) and low coverage of other cell types of interest. To counteract these limitations, we optimized a protocol to isolate nuclei directly from the intact periosteum and from the fracture callus to perform single-nucleus RNA sequencing. This protocol is adapted for fresh murine periosteum, fracture callus, and frozen human periosteum. Nuclei are isolated using mechanical extraction combined with fluorescence-based nuclei sorting to obtain high-quality nucleus suspensions. This protocol allows the capture of the full diversity of cell types in the periosteum and fracture environment to better reflect the in vivo tissue composition.
Key features
• Allows the isolation of nuclei with high-quality RNA for transcriptomic analyses.
• Can be adapted to be used on fresh and frozen tissue.
• Optimized for human and murine periosteum.
Keywords: Nuclei isolation Single-nucleus RNA-seq Periosteum Fracture callus Fluorescence-activated nuclei sorting
Graphical overview
Background
The periosteum is a thin and heterogeneous tissue covering the outer surface of bone that contains skeletal stem/progenitor cells (SSPCs) essential for bone repair. Following a bone fracture, periosteal SSPCs are activated and differentiate to form cartilage and bone cells [1–4]. SSPCs differentiate in a highly complex and dynamic environment marked by the presence of many cell types, including immune cells, endothelial cells, and cells from the nervous system [5,6]. To decipher the heterogeneity of the periosteum and the response of these different cell types to bone fracture, we aimed to perform single nucleus/cell transcriptomics from the periosteum and callus tissues. Cell isolation using enzymatic digestion is commonly used, but there are several disadvantages to this method. First, enzymatic tissue digestion can favor the representation of certain cell types. This usually leads to the overrepresentation of cell types loosely attached to the matrix, such as immune cells, and to the limited coverage of cell types embedded in the matrix, including osteoblasts, osteoclasts, and Schwann cells. Second, FACS-sorting and single cell transcriptomic techniques have constraints in terms of the cell size that can be processed. The fracture callus contains large cells such as hypertrophic chondrocytes and osteoclasts that are not compatible with these techniques. Third, enzymatic digestion needed for single-cell RNA sequencing (scRNAseq) can only be performed on fresh tissues and induces a stress response leading to transcriptomic changes[8]. To overcome these limitations, we optimized and tested a protocol to isolate nuclei from fresh and frozen tissues to perform single-nucleus RNA sequencing (snRNAseq). Several studies have shown that snRNAseq can produce similar results as scRNAseq and capture a greater diversity of cell types [7–11]. Here, we detail three distinct ways of using our protocol in order to isolate nuclei from (i) fresh murine periosteum, (ii) fresh murine fracture hematoma/callus, and (iii) frozen human periosteum. While tissue preparation can change between the different types of samples, the nuclei extraction protocol is common and based on mechanical cell lysis. Nuclei extraction is followed by fluorescence-activated nuclei sorting to eliminate the yields of cellular and matrix debris in the suspension. Overall, this protocol is fast and easy-to-use and is currently the most adequate method to capture the periosteal heterogeneity in vivo.
Materials and reagents
Biological materials
1. 8–12-week-old mice, C57BL6 background (Janvier Labs, France, or equivalent vendors). We used a mix of males and females in our experiments
2. Periosteum from patients. We obtained fresh samples from patients undergoing surgery and processed them up to 4 h after tissue collection. Sample collection from patients requires approval by the Ethics Committee and formal consent from the donor
Reagents
1. DMEM medium (Life Technologies, catalog number: 11966025)
2. HEPES buffer (Thermo Fisher, catalog number: 15630056)
3. Penicillin/streptomycin (pen/strep) (Life Technologies, catalog number: 15140122)
4. PBS, RNase free (Thermo Fisher, catalog number: AM9624)
5. Ethanol absolute (VWR, catalog number: 20821.365)
6. Nuclei lysis buffer (Sigma-Aldrich, catalog number: NUC101-1KT)
7. Bovine serum albumin, molecular biology grade (Merck, catalog number: B6917)
8. RNase inhibitor (Roche, catalog number: 3335399001)
9. DNase/RNase-free distilled water, UltraPure (Life Technologies, catalog number: 10977049)
10. SYTOXTM AADvancedTM Dead Cell Stain kit (Thermo Fisher, catalog number: S10349)
11. DAPI (Life Technologies, catalog number: D3571), resuspend in distilled water at a concentration of 5 mg/mL
12. Buprenorphine (Centravet, catalog number: BUP001)
13. Atipamezole (Centravet, catalog number: ANT201)
14. Ketamine (Centravet, catalog number: KET205)
15. Medetomidine (Centravet, catalog number: DOM003)
Solutions
1. Human sample collection medium (see Recipes)
2. 70% ethanol (see Recipes)
3. Nuclei suspension buffer (see Recipes)
4. DAPI buffer (see Recipes)
Recipes
1. Human sample collection medium
Solution can be stored for up to 1 year at 4 °C.
Reagent Final concentration Volume
DMEM 1× 44.5 mL
HEPES 10% 5 mL
Pen/strep 1% 0.5 mL
Total n/a 50 mL
2. 70% ethanol
Reagent Final concentration Volume
Ethanol (absolute) 70% 700 mL
H2O n/a 300 mL
Total n/a 1,000 mL
3. Nuclei suspension buffer
Make fresh for each experiment and keep at 4 °C on ice.
Reagent Final concentration Amount
RNase-free PBS (10×) 1× 0.5 mL
Bovine serum albumin 2% 0.1 g
RNase inhibitor (40 U/µL) 0.2 U/µL 25 µL
RNase-free water n/a 4.5 mL
Total n/a 5 mL
4. DAPI buffer
Make fresh for each experiment and protect from the light.
Reagent Final concentration Volume
Nuclei suspension buffer 1× 99 µL
DAPI 1/1,000 1 µL
Total n/a 100 µL
Laboratory supplies
1. Conical tubes, 15 and 50 mL (Falcon, catalog numbers: 352097 and 352070 or equivalent)
2. Eppendorf tubes 1.5 and 0.2 mL
3. Cryotube ClearLine® 2 mL (Dutscher, catalog number: 390701)
4. Pipettes 10 and 25 mL (Dutscher, catalog numbers: 357551 and 357535 or equivalent)
5. Pipette tips 1 mL, 200 µL, 20 µL, and 10 µL
6. Falcon 5 mL round-bottom polystyrene tubes (Corning, catalog number: 352235)
7. SterilinTM Quickstart universal containers, PS, 30 mL (VWR, catalog number: 128AR/IRR, or equivalent)
8. Sterile scalpels (Dutscher, catalog number: 132622)
9. Cell strained 40 µm and 100 µm (Fisher Scientific, catalog numbers: 352340 and 352360)
10. 25 G needles (Terumo, catalog number: AN*2516R1)
11. 1 mL syringes (Terumo, catalog number: SS+01H1)
12. Greiner Bio-One Petri dishes (bacterial dish) (Dutscher, catalog number: 633185)
13. Kova® slides (Fisher Scientific, catalog number: 22-270141)
14. Liquid nitrogen
15. Ice
Equipment
1. Centrifuge with temperature control for Falcon 50 mL tubes
2. Centrifuge with temperature control for Eppendorf 1.5 mL tubes
3. BD Influx Cell Sorter or equivalent
4. Zeiss Imager D1 AX10 light microscope (Carl Zeiss Microscopy), or equivalent
5. -80 °C freezer
6. Sterile hood
7. Microvolume pipettes
8. Surgical forceps (Dumont AA Forceps) (FST, catalog number: 11210-20 or equivalent)
9. Surgical scissors (Fine Scissors-ToughCut® 11 mm) (FST, catalog number: 14058-11 or equivalent)
10. Dissecting chisel (Fine Science Tools, catalog number: 10095-12)
11. Drill (Dremel, catalog number: 8050-15)
12. Drill bits (0.4 mm)
13. Heating pad (Harvard Apparatus, catalog number: 55-7033)
14. Trimmer (Kerbl, catalog number: GT416)
15. 15 mL Dounce homogenizer with pestle A (loose) and pestle B (tight) (Sigma-Aldrich, catalog number: D9938)
16. Liquid nitrogen container
17. Ice container
Procedure
A. Prepare material and reagents
1. Set the centrifuge to 4 °C and allow it to cool before use.
2. Prepare all solutions needed for the protocol.
Note: Prepare all reagents in RNase-free conditions. All solutions should be prepared fresh and kept on ice.
Note: The protocol includes two solutions for nuclei processing. The first solution is the nuclei lysis buffer, used to induce cell lysis. The second solution is the nuclei suspension buffer (see Recipes) and corresponds to the buffer used for centrifugation, sorting, and counting of nuclei.
3. Prepare and annotate all tubes needed for the procedure.
Note: To improve the quality of the RNA preparation, it is recommended to work in RNase-free conditions, by using RNase-free solutions and RNase-free materials (tubes, dishes, dissection tools, douncer), and working on clean RNase-free surfaces (use appropriate reagent to clean benches).
B. Option I: Nuclei isolation from fresh murine periosteum (Timing: up to 30 min)
1. Sacrifice the mice by cervical dislocation (or any other appropriate method).
Note: As the periosteum is very thin in adult mice, we recommend using the tibias of at least five mice.
Note: All procedures involving animals must be approved by ethical committees.
2. Rinse the limbs with 70% ethanol.
3. Incise the skin and remove it entirely from the lower limbs. Disconnect the tibias from the limbs by cutting at the knee and ankle level. Place the tibias in a 100 mm sterile Petri dish with RNase-free chilled PBS on ice (Figure 1, left).
Figure 1. Tissue processing of uninjured murine periosteum
4. Remove the soft tissues surrounding the tibias using forceps and scissors (Figure 1, middle left).
Note: Remove soft tissue gently using scissors, as pulling out muscle could detach the periosteum from the cortex.
Note: Dissection steps can be performed under a binocular microscope to reduce the risk of contamination by surrounding tissue.
5. Cut the epiphyses of the tibia using scissors. Flush the bone marrow using RNase-free ice-cold PBS.
6. Collect the periosteum by scraping it from the cortex using a dissecting chisel. Collect the tissue in a 1.5 mL Eppendorf containing ice-cold lysis buffer and place it on ice (Figure 1, middle right).
7. Put the tissues in a 100 mm sterile Petri dish with a drop of ice-cold lysis buffer. Chop it with clean scissors until very small pieces are obtained (Figure 1, right).
8. Proceed immediately to nuclei extraction (section E).
Critical: Do not wait to perform nuclei extraction; any time lost will impact RNA quality.
C. Option II: Isolation of fresh murine fracture hematoma/callus (Timing: up to 30 min)
1. Induce tibial fracture.
Note: All procedures involving animals must be approved by ethical committees.
Note: We recommend using at least five mice for post-fracture day 1 and at least three mice for later time points to obtain a sufficient number of nuclei and overcome interindividual variability.
a. Anesthetize the mice with an intraperitoneal injection of 50 mg/kg ketamine and 1 mg/kg medetomidine. Inject 0.1 mg/kg of buprenorphine subcutaneously for analgesia.
b. After 15–30 min, if the quality of the anesthesia and analgesia is sufficient, shave the right limb and sanitize using skin disinfectant.
Note: The efficiency of anesthesia must be checked using foot pinching or other appropriate techniques.
c. Perform a 2 cm incision on the skin along the tibia using a sterile scalpel and expose the tibial surface by gently separating the muscles from the bone surface.
d. Create three holes in the mid-diaphysis aligned perpendicular to the tibial axis using a drill and a 0.4 mm drill bit.
e. Induce osteotomy by cutting the bone along the three holes with scissors (Figure 2, left).
Figure 2. Tissue processing of murine fracture callus and activated periosteum
f. Close the skin wound using suture threads.
g. Revive the mice with an intraperitoneal injection of 1 mg/kg atipamezole and place it on a 37 °C heating pad until revived.
h. Perform two additional subcutaneous injections of buprenorphine 0.1 mg/kg at 12 and 24 h post-surgery and monitor the mice closely until callus tissue collection.
2. Collect tissue for nuclei extraction.
Note: We tested our protocol on fracture tissue up to 7 days post-fracture. Later time points may need further optimization as the fracture callus becomes more ossified.
a. Sacrifice the mice by cervical dislocation (or any other appropriate method) and rinse the limbs with 70% ethanol.
b. Incise the skin and remove it entirely from the fractured limb. Disconnect the tibia from the limb by cutting at the knee and ankle level and place it in a 100 mm sterile Petri dish with RNase-free chilled PBS.
Note: Proceed very gently to avoid separating the two segments of the tibia while dissecting, as the fractured tibia is very fragile in the first days post-injury.
3. Remove the soft tissues surrounding the tibia using forceps and scissors (Figure 2, middle left).
Note: Remove the soft tissue gently using scissors, as pulling out muscle could damage the fracture tissue.
4. Scrape the fracture hematoma/callus and the activated periosteum from the diaphysis using a dissecting chisel. Collect the tissue in ice-cold lysis buffer placed on ice (Figure 2, middle right).
5. Put the tissue with a drop of ice-cold lysis buffer. Chop it with clean scissors until very small pieces are obtained (Figure 2, right).
6. Proceed immediately to nuclei extraction (section E).
Critical: Do not wait to perform nuclei extraction; any time lost will impact RNA quality.
D. Option III: Isolation of frozen human periosteum
1. Collect and freeze the human periosteum.
a. Place the sample in a tube containing at least 10 mL of ice-cold human sample collection medium immediately after collection. Place at 4 °C until processing (Figure 3, left).
Note: Samples should be processed as early as possible after collection. In our experience, samples can be processed up to 3 h after collection and used for nuclei extraction.
Figure 3. Collection and freezing of the human periosteum
b. Under a cell culture hood, remove the tissue from the tube and place it in a sterile Petri dish. Wash the tissue with RNase-free ice-cold PBS to remove the remaining medium and blood (Figure 3, middle left).
c. Using scissors, scalpel, and tweezers, remove any muscle, fat tissue, or damaged tissue.
d. Cut the periosteum into pieces of at least 0.5 × 0.5 cm. Place the pieces briefly in a dry Petri dish to remove the medium from the tissue (Figure 3, middle right).
Note: We recommend storing larger pieces of tissue to obtain a higher number of nuclei after nuclei extraction. Store at least three pieces of tissues if possible.
e. Place each piece of periosteum in a properly labeled freezing tube.
f. Place the tubes directly in liquid nitrogen and leave for at least 5 min. Then, place the tubes at -80 °C (Figure 3, right).
Caution: Handle liquid nitrogen with care and follow safety rules.
Stop point: Samples can be stored at -80 °C for a long time before processing. We have used samples for up to three years after freezing.
2. Check RNA integrity.
a. If possible, use one sample from the same batch to extract RNA and check that RNA integrity is above 6.5. We routinely used the RNeasy Mini Kit following the manufacturer’s instructions for RNA extraction followed by RIN measure on Agilent TapeStation, but any equivalent methods can be used.
Note: We avoid using samples with RIN lower than 6.5 as it may lead to poor RNA quality after nuclei extraction.
3. Prepare tissue for nuclei extraction (Timing: up to 15 min).
a. Take the sample from the -80 °C freezer and put it on dry ice (Figure 4, left).
Critical point: Keep the tube on dry ice until it is used for extraction. Do not allow the sample to thaw before placing it into the lysis buffer.
b. Put the frozen tissue on an RNase-free Petri dish placed on ice. Add a few drops of ice-cold lysis buffer (Figure 4, middle).
c. Chop the tissue with clean scissors until very small pieces are obtained (Figure 4, right).
d. Proceed immediately to nuclei extraction (section E).
Critical: Do not wait to perform nuclei extraction; any time lost will lead to a decrease in RNA quality.
Figure 4. Tissue processing of human periosteum
E. Nuclei extraction (Timing: up to 15 min)
1. Put the tissues prepared in parts B, C, or D in a glass douncer. Add up to 7 mL of ice-cold lysis buffer (Figure 5, left, middle left).
Figure 5. Nuclei extraction from murine and human periosteum
2. Put the douncer on ice for 2 min.
3. While keeping the douncer on ice, lower and raise the pestle A gently 15–20 times, avoiding bubble formation (Figure 5, middle right).
Note: There shouldn’t be too much resistance while using pestles. Resistance can be due to the presence of big pieces of tissue, which would require better chopping before putting the tissue in the douncer, or to a high ratio of tissue/lysis buffer, which would require putting less tissue or more lysis buffer, if possible.
4. Lower and raise the pestle B gently up to 10 times (Figure 5, right).
5. Filter the suspension with a 100 µm cell strainer in a 50 mL conical tube. Collect the flowthrough and filter using a 40 µm cell strainer in a clean 50 mL tube.
6. Centrifuge at 500× g for 5 min at 4 °C. Carefully remove the supernatant without disturbing the pellet.
7. Add 1 mL of ice-cold nuclei suspension buffer. Resuspend the pellet and transfer to an RNase-free Eppendorf tube.
8. Centrifuge at 500× g for 5 min at 4 °C. Carefully remove the supernatant without disturbing the pellet.
9. Resuspend in 200 µL of ice-cold nuclei suspension buffer. Proceed to nuclei sorting.
F. Nuclei sorting and counting (Timing: up to 30 min)
1. Nuclei sorting:
a. Add 1 µL of SYTOXTM AADvancedTM in the solution to label DNA in nuclei.
Note: We also tested our protocol using DAPI staining with similar results.
b. Sort up to 150,000 nuclei (Sytox AADvanced+ nuclei) using the appropriate gating strategy (Figure 6). Collect in 1.5 mL Eppendorf tubes containing 0.75 mL of ice-cold nuclei suspension buffer.
Note: Prior to the snRNAseq experiment, cell sorting should be settled with appropriate controls including unstained nuclei.
c. Centrifuge the suspension of sorted nuclei at 500× g for 5 min at 4 °C. Carefully remove the supernatant without disturbing the pellet.
Critical: Depending on the number of sorted nuclei, the pellet may not be visible. Proceed carefully to the supernatant removal to avoid eliminating the nuclei.
d. Resuspend in 50 µL of ice-cold nuclei suspension buffer.
Figure 6. Sorting strategy to obtain a nuclei suspension without debris. The suspension is first gated based on SSC-A and FSC-A to remove larger debris (left), and nuclei are selected based on DAPI fluorescence (right).
2. Nuclei counting (Timing: up to 15 min):
a. Mix 5 µL of the nuclei suspension with 10 µL of DAPI solution in a 0.2 mL tube.
Note: Adapt the dilution to obtain an optimal number of nuclei for counting. Too much or not enough nuclei during counting could lead to errors.
b. Transfer 10 µL of the stained nucleus suspension to a Kova slide.
c. Under a fluorescence microscope, check the quality of the nuclei suspension. Count the number of nuclei to obtain the concentration of the nucleus suspension (Figure 7).
Note: We recommend using both brightfield and fluorescence microscopy to check the nuclei suspension.
Critical: It is crucial to assess the quality of the nuclei suspension before proceeding to the next steps of the experiments. The suspension should only contain nuclei and no debris. The shape of the nuclei reflects their quality. If the suspension contains debris or a high percentage of low-quality nuclei, we recommend not to use it for further analysis.
Figure 7. Nuclei counting and quality check. Nuclei should have a clean round shape. Nuclei with abnormal shape, unclear boundaries, and altered membranes are considered low quality.
3. Proceed immediately to loading for snRNAseq following the manufacturer’s instructions.
Note: Using a nuclei solution of 700–1,200 nuclei per microliter is recommended.
Validation of protocol
This protocol was adapted from previously published protocols [12,13] and was used in other studies and on various tissues. We have generated more than 10 murine samples and 6 human samples using this protocol, which were published in [3,5,6]. Datasets from the murine uninjured periosteum and fracture callus are available at the following link: https://cells.ucsc.edu/?ds=fracture-repair-atlas.
General notes and troubleshooting
Troubleshooting
Poor data quality after snRNAseq might be due to low-quality nuclei preparation. To improve sample quality, we recommend the following:
1. Work in an RNase-free environment. All solutions used for the protocol must be RNase-free. All tools used for this protocol must be sterile and cleaned to eliminate RNase.
2. Reduce experimental time. Time is crucial to preserve RNA quality. Optimize all steps of nuclei extraction and sorting and process immediately to snRNAseq after nuclei extraction.
3. A poor yield of nuclei can be due to problems in mechanical extraction. The use of the pestle should be smooth. If there is strong resistance while using the pestle, consider increasing the chopping time, reducing the amount of tissue, or increasing the volume of lysis buffer.
4. Ensure that the nuclei counting is accurate and that the suspension does not contain debris.
5. If working with frozen human periosteum, proper snap freezing and cryopreservation are required to obtain high-quality nuclei and RNA. Reduce the time between tissue collection and freezing. To ensure RNA quality, perform RNA extraction and check RNA integrity (RIN) from a sample from the same batch. Samples with a RIN lower than 6.5 will lead to poor results.
6. Differences in age, sex, species, and pathological conditions can vary the yield of recovered nuclei and should be taken into consideration. Adjust the amount of tissue for extraction to compensate.
Acknowledgments
This work was supported by ANR-21-CE18-007-01, NIAMS R01 AR081671, and US Department of the Army NF220019 to C.C. S. Perrin, M. Ethel, and C. Goachet were supported by a PhD fellowship from Paris Cité University, Univ Paris-Est Creteil, and Fondation pour la Recherche Médicale, respectively. This protocol was first described by Perrin et al. [3] and Perrin et al. [6].
Competing interests
Authors declare no competing interest.
Ethical considerations
All procedures involving animals were approved by the Paris Est Creteil University Ethical Committee (agreement #19295-2019052015468705). Sample collection from patients was approved by the Ethics Committee CPP-IDF-2 (#ID-RCB/EUDRACT: 2014-A01420-47; IMNIS2014-03).
References
Duchamp de Lageneste, O., Julien, A., Abou-Khalil, R., Frangi, G., Carvalho, C., Cagnard, N., Cordier, C., Conway, S. J. and Colnot, C. (2018). Periosteum contains skeletal stem cells with high bone regenerative potential controlled by Periostin. Nat Commun. 9(1): 773.
Jeffery, E. C., Mann, T. L., Pool, J. A., Zhao, Z. and Morrison, S. J. (2022). Bone marrow and periosteal skeletal stem/progenitor cells make distinct contributions to bone maintenance and repair. Cell Stem Cell. 29(11): 1547–1561.e6.
Perrin, S., Ethel, M., Bretegnier, V., Goachet, C., Wotawa, C. A., Luka, M., Coulpier, F., Masson, C., Ménager, M., Colnot, C., et al. (2024). Single-nucleus transcriptomics reveal the differentiation trajectories of periosteal skeletal/stem progenitor cells in bone regeneration. eLife. 13: e92519.
Perrin, S. and Colnot, C. (2022). Periosteal Skeletal Stem and Progenitor Cells in Bone Regeneration. Curr Osteoporos Rep. 20(5): 334–343.
Hachemi, Y., Perrin, S., Ethel, M., Julien, A., Vettese, J., Geisler, B., Göritz, C. and Colnot, C. (2024). Multimodal analyses of immune cells during bone repair identify macrophages as a therapeutic target in musculoskeletal trauma. Bone Res. 12(1), 56.
Perrin, S., Protic, S., Bretegnier, V., Laurendeau, I., de Lageneste, O. D., Panara, N., Ruckebusch, O., Luka, M., Masson, C., Maillard, T., et al. (2024). MEK-SHP2 inhibition prevents tibial pseudarthrosis caused by NF1 loss in Schwann cells and skeletal stem/progenitor cells. Sci Transl Med. 16(753): eadj1597.
Ding, J., Adiconis, X., Simmons, S. K., Kowalczyk, M. S., Hession, C. C., Marjanovic, N. D., Hughes, T. K., Wadsworth, M. H., Burks, T., Nguyen, L. T., et al. (2020). Systematic comparison of single-cell and single-nucleus RNA-sequencing methods. Nat Biotechnol. 38(6): 737–746.
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Wen, F., Tang, X., Xu, L. and Qu, H. (2022). Comparison of single‑nucleus and single‑cell transcriptomes in hepatocellular carcinoma tissue. Mol Med Rep. 26(5): e12855.
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Article Information
Publication history
Received: Oct 17, 2024
Accepted: Dec 29, 2024
Available online: Jan 22, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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5,212 | https://bio-protocol.org/en/bpdetail?id=5212&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
A Bio-protocol resource
This is an In Press version of the protocol that has not yet been assigned to an issue.
Peer-reviewed
Voltage clamp fluorometry in Xenopus laevis oocytes to study the voltage-sensing phosphatase
VY Victoria C. Young
Vamseedhar Rayaprolu
SK Susy C. Kohout
In Press, Available online: Jan 19, 2025
DOI: 10.21769/BioProtoc.5212 Views: 23
Reviewed by: Willy R Carrasquel-UrsulaezAkira KarasawaXiaochen SunVinaykumar Idikuda
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Original Research Article:
The authors used this protocol in Journal of General Physiology Jul 2024
Abstract
Voltage clamp fluorometry (VCF) is a powerful technique in which the voltage of a cell’s membrane is clamped to control voltage-sensitive membrane proteins while simultaneously measuring fluorescent signals from a protein of interest. By combining fluorescence measurements with electrophysiology, VCF provides real-time measurement of a protein’s motions, which gives insight into its function. This protocol describes the use of VCF to study a membrane protein, the voltage-sensing phosphatase (VSP). VSP is a 3 and 5 phosphatidylinositol phosphate (PIP) phosphatase coupled to a voltage sensing domain (VSD). The VSD of VSP is homologous to the VSD of ion channels, with four transmembrane helices (S1–S4). The S4 contains the gating charge arginine residues that sense the membrane’s electric field. Membrane depolarization moves the S4 into a state that activates the cytosolic phosphatase domain. To monitor the movement of S4, the environmentally sensitive fluorophore tetramethylrhodamine-6-maleimide (TMRM) is attached extracellularly to the S3-S4 loop. Using VCF, the resulting fluorescence signals from the S4 movement measure the kinetics of activation and repolarization, as well as the voltage dependence of the VSD. This protocol details the steps to express VSP in Xenopus laevis oocytes and then acquire and analyze the resulting VCF data. VCF is advantageous as it provides voltage control of VSP in a native membrane while quantitatively assessing the functional properties of the VSD.
Key features
• Voltage clamp fluorometry using Xenopus laevis oocytes expressing the voltage-sensing phosphatase of Ciona intestinalis.
• This protocol uses the fluorophore tetramethylrhodamine-6-maleimide (TMRM).
• This protocol details the procedure for a two-electrode voltage clamp using the Dagan CA-1B amplifier.
Keywords: Voltage clamp fluorometry (VCF) Two-electrode voltage clamp (TEVC) Tetramethylrhodamine-6-maleimide (TMRM) Voltage-sensing phosphatase (VSP)
Graphical overview
Background
Voltage clamp fluorometry (VCF) is a powerful technique to study voltage-sensitive conformational dynamics in membrane proteins. By combining the membrane voltage clamp with the measurement of fluorescent signals from the fluorophore attached to the protein of interest, real-time measurements of the protein’s conformational changes can be obtained and correlated to the electrophysiological functions. Initially developed to measure the conformational changes occurring in ion channels [1,2], the technique evolved to study other membrane proteins such as ion transporters [3,4] and, as discussed in this protocol, the voltage-sensing phosphatase (VSP) [5,6].
VSP is a 3 and 5 phosphatidylinositol phosphate (PIP) phosphatase coupled to a voltage-sensing domain (VSD) [7]. The VSD is made up of four transmembrane helices (S1–S4), with the S4 containing arginine residues that act as gating charges to sense the electric field of the membrane. Depolarization of the membrane causes the S4 to move from a down “resting” state in the membrane to an up “active” state. Unlike ion channels, the movement of the VSD in VSP does not lead to ionic currents; therefore, VCF is particularly advantageous as a robust measure of VSP’s voltage sensitivity and protein motions.
Environmentally sensitive fluorophores, like tetramethylrhodamine-6-maleimide (TMRM), are often used to monitor protein movements when attached to a protein’s predicted mobile region. Because the maleimide of TMRM is thiol-reactive, single cysteine mutations are introduced into these regions for labeling and are empirically tested for fluorescence changes that correlate with protein activation. To monitor S4 movements in VSP, G214C, at the top of S4 in the extracellular loop between S3 and S4, is a common labeling site. By analyzing the G214C-TMRM fluorescence signal during activation and repolarization conditions, the kinetics of the VSD, specifically S4, are calculated, giving us insight into the VSD motions in response to the changing voltage stimulus. Additionally, by taking the amplitude of the fluorescence signal relative to the membrane voltage, we calculate the voltage dependence of the VSD, which correlates with the voltage dependence of VSP’s phosphatase activity. Further, VCF labeling sites can be combined with other mutations to probe the mechanism of voltage dependence [6,8], coupling properties of the linker [9–11] and phosphatase activity [12,13]. While we only discuss the G214C labeling site here, different labeling sites will report on different protein motions and can be used to further test the mechanism of protein function [10,11].
Because Xenopus laevis (X. laevis) oocytes are large single cells that are easy to manipulate and maintain, they are a great expression system for VCF. This protocol describes all the different parts of preparing, setting up, and conducting a VCF experiment. It starts with how to prepare X. laevis oocytes from the ovary; it then describes how to make the messenger RNA (mRNA). It moves on to how to inject the oocytes with the mRNA for expression, how to label them with TMRM, and ultimately how to voltage clamp them using a Dagan CA-1B amplifier for two-electrode voltage clamp. Lastly, this protocol describes the steps for analyzing the acquired fluorescence signals.
Materials and reagents
Biological materials
1. Xenopus laevis ovaries (Xenopus One, catalog number: 10004, ¼ ovary)
2. Ciona intestinalis voltage-sensing phosphatase (Ci-VSP) in pSD64TF vector (Y. Okamura, Osaka University, Osaka, Japan, Addgene plasmid #80332) (Figure 1A)
Figure 1. Map of pSD64TF Ci-VSP plasmid. Vector of pSD64TF with ampicillin selection and an SP6 promoter. The VSD, catalytic domain, and C2 domain of Ci-VSP are noted in the gray regions.
Reagents
1. mMESSAGE mMACHINE SP6 Transcription kit (Invitrogen, catalog number: AM1340)
2. Linearization enzyme XbaI (NEB, catalog number: R3131L)
3. Water, DNase/RNase free (Fisher, catalog number: BP561-1)
4. Ethyl alcohol 200 proof (EtOH) (Pharmco, catalog number: 111000200)
5. Ethyl alcohol 190 proof (EtOH) (Pharmco, catalog number: 111000190)
6. GeneRuler DNA ladder (Thermo Scientific, catalog number: SM0333)
7. Millennium RNA marker (Thermo Fisher, catalog number: AM7150)
8. QIAquick PCR Purification kit (QIAGEN, catalog number: 28104)
9. Light mineral oil (Millipore Sigma, catalog number: ES005C)
10. Collagenase type 2 (Worthington Biochemical Corporation, catalog number: LS004177)
11. Collagenase type 3 (Worthington Biochemical Corporation, catalog number: LS004183)
12. Tetramethylrhodamine-6-maleimide (TMRM) (Abcam, catalog number: ab145471)
13. Sodium pyruvate (Fisher, catalog number: BP356-100)
14. NaCl (Fisher, catalog number: BP358-1)
15. KCl (Fisher, catalog number: BP366-500)
16. CaCl2·2H2O (JT Baker, catalog number: 1332-01)
17. MgCl2·6H2O (Thermo Scientific, catalog number: J62575.36)
18. Gentamicin sulfate (Thermo Scientific, catalog number: J6283406)
19. HEPES (Fisher, catalog number: BP310-500)
20. N,N-Dimethylformamide (DMF) (Thermo Scientific, catalog number: 041859AK)
21. HCl (Fisher, catalog number: S25358)
22. NaOH (Fisher, catalog number: P250-500)
23. RNase Away (Thermo Scientific, catalog number: 7005-11)
Solutions
1. ND-96 (see Recipes)
2. Ca2+-free buffer (see Recipes)
3. ND-96(-) (see Recipes)
4. Bridge buffer (see Recipes)
Recipes
1. ND-96 (1 L)
Note: Adjust pH to 7.6 with HCl or NaOH and filter-sterilize to prevent contaminating growth. Store at room temperature for up to a year or longer as long as it does not show any growth.
Reagent Final concentration Quantity or Volume
NaCl 96 mM 56.1 g
KCl 2 mM 0.15 g
CaCl2·2H2O 1.8 mM 0.265 g
MgCl2·6H2O 1 mM 0.203 g
Gentamicin sulfate 50 μg/mL 2 mL of 50 mg/mL stock
Sodium pyruvate 2.5 mM 5 mL of 1 M stock
HEPES 10 mM 2.383 g
Total n/a 1,000 mL
2. Ca2+-free buffer (1 L)
Note: Adjust pH to 7.6 with HCl or NaOH and autoclave to sterilize. Store at room temperature for up to a year or longer as long as it does not show any growth.
Reagent Final concentration Quantity or Volume
NaCl 83 mM 4.8 g
KCl 2 mM 0.15 g
MgCl2·6H2O 1 mM 0.203 g
HEPES 10 mM 2.383 g
Total n/a 1,000 mL
3. ND-96(-) (1 L)
Note: Adjust pH to 7.6 with HCl or NaOH and autoclave to sterilize. Store at room temperature for up to a year or longer as long as it does not show any growth.
Reagent Final concentration Quantity or Volume
NaCl 96 mM 56.1 g
KCl 2 mM 0.15 g
CaCl2·2H2O 1.8 mM 0.265 g
MgCl2·6H2O 1 mM 0.203 g
HEPES 10 mM 2.383 g
Total n/a 1,000 mL
4. Bridge buffer (1 L)
Note: Adjust pH to 7.4 with HCl or NaOH and autoclave to sterilize. Store at room temperature for up to a year or longer as long as it does not show any growth.
Reagent Final concentration Quantity or Volume
NaCl 1 M 58.4 g
HEPES 10 mM 2.383 g
Total n/a 1,000 mL
Laboratory supplies
1. Reusable 45 mm bottle top filter (Nalgene, catalog number: DS0320-5045)
2. Membrane and prefilter disks (Nalgene, catalog number: DS02154020)
3. Low-retention filtered RNase-free pipette tips (Fisher, catalog numbers: 02-707-002, 02-707-006, and 02-707-008)
4. 3.5” glass capillaries (Drummond Scientific, catalog number: 3-000-203-G/X)
5. Glass capillary tubes (VWR, catalog number: 5432-921)
6. Glass coverslips, No. 1, 22 × 40 mm (Warner Instruments, catalog number: 64-0707)
7. Polypropylene microcentrifuge tubes (Globe Scientific, catalog number: 11563)
8. Falcon Petri dishes (Falcon, catalog number: 351007)
9. 50 mL conical tubes (Thermo Fisher, catalog number: AM12502)
10. 25 G × 5/8 in. needle (BD, catalog number: 305122)
11. 1 mL syringe (BD, catalog number: 309628)
12. DrieriteTM 10–20 mesh (Thermo Scientific, catalog number: 219065000)
13. 150 mm × 15 mm Petri dish (Falcon, catalog number: 351058)
14. Palladium wire (Thermo Scientific, catalog number: AA45072G1)
Equipment
1. Belly Dancer Orbital Shaker (IBI Scientific, model: BDRAA115S)
2. Glass Pasteur pipette (Fisher, catalog number: 13-678-4A)
3. Blunted glass Pasteur pipette (homemade using the pipettes in #2)
4. Mini low-temperature refrigerated incubator, 18 L (Fisher, catalog number: 15-015-2632)
5. Water bath 2 L digital (Benchmark Scientific, catalog number: B2000-2)
6. Refrigerated high-speed microcentrifuge (Thomas Scientific, catalog number: 1154Q52)
7. Rotor (Thomas Scientific, model: AS-24-2)
8. Centrifugal vacuum concentrator (Thermo Scientific, model: DNA120)
9. NanoDropTM One spectrophotometer (Thermo Scientific, catalog number: ND-ONE-W)
10. Nanoject II Auto-Nanoliter Injector (Drummond Scientific, catalog number: 3-000-204)
11. Dissecting microscope (Olympus, model: SZ61)
12. Manipulator (Märzhäuser Wetzlar, catalog number: 00-42-101-0000)
13. Flaming/Brown micropipette puller (Sutter Instruments, model: P-97)
14. Inverted microscope (Leica, model: DM IRBE, catalog number: 020-525.701 to 020-525.780)
15. HC Pl APO 20×/0.7 fluorescence objective (Leica, catalog number: 506166)
16. Amplifier (Dagan Corporation, model: CA-1B)
17. Photomultiplier tube (PMT) (ThorLabs, catalog number: PMTSS2)
18. Axon Digidata-1440A (Molecular Devices Instruments, catalog number: DD1440A)
19. X-Cite XLED1 light source (Lumen Dynamics, catalog number: 010-00288R)
20. LED 505-546 nm (Lumen Dynamics, model: BGX)
21. Cy3 Leica cube set with HQ531/40 excitation filter, HQ593/40 emission filter, Q562LP dichroic (Semrock, catalog number: Cy3-4040C-LSC-ZERO)
22. Eight-pole Bessel filter (Frequency Devices, model: 900CT)
23. Gravity glass puller (Narishige, model: PC-10)
24. Vibration isolation platform (TMC, catalog number: 77049189)
25. Faraday cages (homemade)
26. Glass agarose bridges (homemade)
27. Bath chamber (Warner Instruments, model: RC-24E)
28. Bath chamber platform (Warner Instruments, model: 64-1526)
29. Inox tweezers style #5 (Dumont, catalog number: 11254-20)
30. Mesh 0.011 diameter, 18 × 16 mesh count (Phifer, catalog number: 3002201)
Software and datasets
1. pClamp version 10.3 software package (Axon Instruments)
2. Microsoft Excel, version 16.9 for Mac (Microsoft)
3. Igor Pro version 8 software (WaveMetrics)
Procedure
A. Defolliculating X. laevis oocytes
1. Immediately upon arrival of the X. laevis ovaries (Figure 2A and B), wash the ovary pieces in Ca2+-free buffer.
a. If unable to immediately process the ovaries, store the oocytes in fresh ND-96 with antibiotic in the incubator at 18 °C for a maximum of 4 h. Do NOT freeze or store at 4 °C.
Note: We use ND-96 with and without antibiotic. We use the notation ND-96(-) to denote without antibiotic and just ND-96 to denote with antibiotic.
Figure 2. X. laevis oocytes pre-digestion of follicular membrane. A. Lobe of X. laevis ovary. B. Magnified lobe of X. laevis ovary; black arrow highlights the follicular membrane. C. Morselized ovary pieces.
2. Use sterile tweezers and/or small sharp scissors to morselize the ovary into pieces around 5 mm in size (which is about five oocytes) in Ca2+-free buffer (Figure 2C).
Tip: Make the morselized pieces roughly the same size so the follicular layer of the oocyte digests at the same rate.
3. Place the morselized ovary pieces into a 50 mL tube and wash to remove the yolk of lysed oocytes.
a. Agitate by gently inverting the tube up and down. Pour out the Ca2+-free buffer and replace with fresh Ca2+-free buffer. Repeat the wash steps 2–3 times. On the final wash, decant as much of the liquid off as you can, leaving ~5 mL of morselized ovary pieces.
4. Digest the cleaned and morselized ovary pieces.
a. In a 50 mL tube, make the digestion solution: 0.5 mg/mL collagenase type II and 0.83 mg/mL collagenase type III in 15 mL of Ca2+-free buffer.
b. Add the digestion solution to the ~5 mL of morselized ovary pieces and gently shake on the belly dancer rotating platform at speed ~4 at room temperature for 60–70 min.
Note: Collagenase activity can show batch-to-batch variability. If it is the first use of a collagenase batch, it is recommended to check the progression of digestion earlier in the process (~30 min) and then repeatedly check every 10 min. Once a good baseline of the collagenase activity is established, the amount of monitoring can be reduced. For our batches, checking at the initial 60 min mark is a good starting point to monitor the collagenase effectiveness.
c. Partway through digestion, remove a few oocytes from the digestion solution. Place them onto a Petri dish under a dissecting microscope to assess the level of digestion through the removal of the follicular membrane (Figure 3A). The follicular membrane is part of the extracellular matrix that holds the cells together. On an individual cell, it looks like a transparent layer with red veining (Figure 3C).
Figure 3. X. laevis oocytes post-digestion of the follicular membrane. A. Mixture in the appearance of X. laevis oocytes post-digestion. B. Various qualities of X. laevis oocytes. From left to right: small immature cells, poor-quality stage-V cells, and good-quality stage-V cells. C. Magnified image of a good-quality stage-V cell with part of the follicular membrane attached.
5. After the majority of the morselized ovary is digested into individual oocytes, and approximately 20%–50% have the follicular membrane removed, wash the oocytes in the same 50 mL tube by gently inverting the tube up and down. Pour out the Ca2+-free buffer and replace with 30 mL of fresh Ca2+-free buffer. Repeat this wash step 10 times.
6. Sort the healthy stage-V oocytes from the dead, immature, and unhealthy oocytes. A healthy stage-V oocyte will be approximately 1 mm in diameter with a well-defined line for animal (dark) and vegetal (light) poles. Any oocytes that are small or have mixing animal and vegetal poles are discarded (Figure 3B). Culture the remaining healthy oocytes in ND-96 at 18 °C.
a. To create a glass pipette for moving the oocytes, a glass Pasteur pipette is first scored by a file close to the end of the taper of the pipette. The placement is dependent on how big an opening is desired. The glass is broken off at the scoring and briefly passed through a flame to create a smooth edge to the glass. Allow it to cool to room temperature before use (Figure 4A).
Figure 4. Glass Pasteur pipettes for X. laevis oocyte manipulation. A. Pasteur pipettes for moving oocytes. Top: some pipette taper remaining and a small opening; bottom: pipette taper removed and a larger opening. B. Pasteur pipettes for manual defolliculation. Top: more taper and longer length; bottom: less taper and shorter length.
7. In our experience, oocytes have better experimental survival under voltage clamp at high positive voltages if they are manually defolliculated rather than completely defolliculated by collagenase.
a. For manual defolliculation, select individual oocytes with the follicular membrane still attached (Figure 3C).
b. Pipette the oocytes into a Falcon Petri dish containing ND-96.
c. To create a defolliculating glass pipette, repeat step 6a but heat the end of the glass for longer to create a blunted tip that is smaller than the size of the oocyte. Allow to cool to room temperature before use (Figure 4B).
d. Gently press the folliculated oocytes into the Falcon dish with the blunted glass pipette until the follicular membrane “sticks” to the Falcon dish (Video 1). While the exact chemistry is uncertain, we believe the crystal-grade polystyrene of the Falcon Petri dishes is more hydrophobic than other Petri dishes and therefore allows for better follicular membrane sticking.
e. Using the blunted glass pipette, gently roll the oocyte out of the stuck follicular membrane.
f. Repeat for all the oocytes needed. We typically inject approximately 10–20 oocytes.
Tip: We find that lining up all the oocytes in rows in the dish facilitates this process.
Video 1. Manual defolliculation of an oocyte. The glass probe approaches from the left and gently presses the oocyte into the Falcon dish until the follicular membrane sticks. The probe is then used to gently roll the oocyte away.
B. In vitro RNA production
1. Critical: The following steps should be done under DNase/RNase-free conditions. Use DNase/RNase-free plasticware and reagents. Wear gloves and clean with RNase decontamination solution such as RNase Away in the following order: benchtop, pipettors, and gloves.
2. Linearize the Ci-VSP plasmid in the vector of pSD64TF.
a. Set up the reaction mixture to contain 5 μg of Ci-VSP plasmid, 5 μL of the restriction enzyme Xba I, and 10 μL of buffer rCutSmartTM, and bring to a total reaction volume of 100 μL with nuclease-free water.
b. Incubate the reaction in a water bath incubator at 37 °C for 1 h.
3. Clean up the linearization reactions with the QIAquick PCR Purification kit using RNase-free plasticware and H2O. Follow the kit instructions and then elute in 30 μL of RNase-free water supplied in the transcription kit.
a. Afterward, run 0.5–1 μL of the elution on a DNA gel to check for complete linearization. Complete linearization is verified by the presence of a single band at the molecular weight of the plasmid (~5 kb, Figure 5A, right lane). The circular plasmid will run further (Figure 5A, middle lane) due to the supercoil conformation.
Figure 5. DNA and RNA gels. A. Lanes from left to right: GeneRuler DNA ladder, Ci-VSP circular plasmid, Ci-VSP linearized DNA. B. Left lane: MillenniumTM Marker RNA ladder; right lane: Ci-VSP mRNA product.
4. Transcribe RNA with the mMESSAGE mMACHINE SP6 Transcription kit.
a. Set up the reaction to contain 1 μg of linearized and purified DNA (6 μL), 2 μL of reaction buffer, 10 μL of nucleotide mixture, and 2 μL of polymerase enzyme in a total reaction volume of 20 μL. Doubling the reaction also works well.
b. Incubate this reaction in a water bath incubator at 37 °C for 2–4 h.
Tip: We recommend incubating the reaction for the full 4 h to obtain the maximum amount of RNA.
5. Make the LiCl precipitation solution by adding 30 μL of nuclease-free H2O to 20 μL of LiCl from the RNA transcription kit (7.5 M LiCl, 50 mM EDTA). If the reaction is doubled, then the precipitation solution should also be doubled.
a. Add the LiCl precipitation solution to the RNA reaction.
b. Incubate at -20 °C for a duration of 20 min to overnight. We tend to get better precipitation with longer incubations.
6. Spin the stopped reaction at max speed (≥17,000× g) for 30 min at 4 °C. A white pellet may be visible here, but in many cases, it may be clear.
Tip: Spin the tube with the square latch side up. This will pellet the RNA against the wall of the square latch so that even when the pellet is poorly visible, care can be taken around this area of the tube to prevent accidentally pipetting away the pellet in the following step. As the RNA pellet becomes more visible throughout the wash steps, keep track of the pellet.
7. Carefully pipette off the supernatant so as to not disturb the RNA pellet and wash with 500 μL of cold RNase-free 70% EtOH.
a. Spin the tube at max speed (≥17,000× g) for 5 min at 4 °C.
Note: RNase-free 200 proof EtOH is diluted to 70% using RNase-free H2O and stored at -20 °C.
8. Remove 70% EtOH and add fresh 70% EtOH. Repeat step B7 for a total of three washes. Keep the sample cold at all times during the washes. An ice bucket is sufficient.
9. After the last 70% EtOH wash, carefully and completely pipette off the EtOH and dry the RNA pellet at a medium heat setting in a vacuum concentrator for 2 min. You can also leave the tubes on a bench with the lids open to allow for evaporation; however, we recommend the vacuum concentrator to limit RNase contamination from leaving the lids open.
10. Resuspend the dried RNA pellet in 10–20 μL of nuclease-free H2O. If the RNA pellet is large, heat at 70 °C for 2 min to completely dissolve the RNA pellet.
a. RNA is stored at -20 °C in its original resuspended concentration. While we have not seen issues with freeze/thaw degradation, 2–5 μL aliquots can be made. In our hands, stock RNA stored at -20 °C can last for several years even at stock concentrations. RNA can also be stored at -80 °C and can last a decade or longer.
11. Run 0.5 µL of the final RNA product on an RNA gel to check for the quality of the product.
Note: Remove any secondary RNA structure by heating the RNA mixed with the formaldehyde loading buffer that comes with the transcription kit to 70 °C for 10 min and then cooling on ice for 2 min before loading onto the gel. Not all RNA will form a secondary structure, but the Ci-VSP RNA does.
a. Look for bands at the expected molecular weight between 3,000 and 2,000 bp (Figure 5B).
12. To assess the quantity and quality of the final RNA product, measure on the NanoDropTM. A260/A280 should be ~2. Use RNase-free water as blank. Concentrations will depend on how much water is used to resuspend the pellet. We typically get between 1 and 2 µg/µL.
13. Before injection, dilute Ci-VSP mRNA to 0.4–0.8 µg/µL depending on the expected incubation time. The higher the amount of injected mRNA, the shorter the time needed for incubation.
a. Measure again on the NanoDropTM to ensure the accuracy of the dilution.
C. RNA injection
1. Critical: These steps should be done under RNase-free conditions as discussed in section B.
2. Prepare the injection glass.
a. Pull 3.5” glass capillaries with the P-97 Sutter Instruments puller.
i. Each puller’s filament is different, so per the instruction manual, determine the ramp number for the glass being used. The ramp value for our filament is 561.
ii. Use the ramp value to create a program that will pull the glass into a long needle-like projection (Figure 6A) (see the Pipette Cookbook from Sutter Instruments).
Tip: We pull a maximum of four glass injection needles before turning the puller off to allow it to cool for at least 15 min before pulling another four glass injection needles (if needed). This pause period helps maintain the quality of the injection glass needles pulled by not overheating the filament.
Figure 6. Supplies and setup for X. laevis injection. A. Glass injection needles. Top: directly from the glass pipette puller; bottom: after being trimmed. B. Microscope view of the needle aligned with the parafilm corner and the mineral oil being ejected out. C. View of the injector and injection needle with mineral oil being ejected out. The metal plunger is halfway down the needle. D. Oocytes resting in the etched Petri dish.
b. Break the tip of this injection glass with tweezers to give a suitable opening for injection (Video 2). The opening diameter we usually use is approximately 45 µm. The smaller the diameter, the less damage is done to the oocyte membrane; however, the more likely the injection needle is to get clogged (Figure 6A).
Tip: The glass tip coming out of the puller should be thin enough to be flexible. We aim our tweezers to the area where the glass bends and then break just below that region.
c. Backfill the glass needle with 100% mineral oil using a 25-gauge needle and syringe.
Tip: Take care not to produce any air bubbles in the mineral oil as this will negatively impact the injection volume’s accuracy.
d. On the Nanoject II injector, loosen the black collet and carefully slide the injection glass needle onto the metal plunger pushing until the glass hits the white spacer (hidden under the black collet) and then tighten the collet. Double-check that the glass needle is secure on the injector by giving a gentle tug.
e. Cut a square of parafilm and apply 70% EtOH (made from 190 proof EtOH) to the work surface to act as a lubricant for moving the parafilm around. Place the parafilm on the work surface with the paper side up and remove the paper. This is assumed to be the RNase-free side.
f. Line up a corner of the parafilm in the view of the microscope; this now becomes the point toward which to aim the tip of the needle. With the needle touching the parafilm, press the empty button on the injector until the metal plunger reaches approximately halfway down the glass needle. Then, pull the needle directly up from the parafilm (Figure 6B and C).
i. Another method is to extend the plunger before placing the needle on the injector. This avoids expelling a significant amount of mineral oil but requires care not to bend the plunger when placing the needle. Both methods work equally well.
g. Place 0.5–1 μL of diluted RNA (depending on how many cells you want to inject at once) onto the piece of parafilm. Find the drop of RNA in the microscope by moving the parafilm around. Then, lower the glass needle into the center of the RNA drop, making sure no air is present in the tip, and press the fill button to fill the needle with RNA.
Tip: Sometimes, letting a drop of oil cover the RNA makes this step easier since the RNA will not evaporate as quickly. DO NOT fill the needle with the oil once the RNA is gone.
3. Place the defolliculated oocytes into a solution of ND-96 on an etched plastic Petri dish to prevent rolling (Figure 6D).
4. At a 45° angle, push the glass needle into the oocyte until it just slightly punctures the membrane. There is no need to go deep into the cell. We recommend creating a divot on the cell membrane first and then pushing in with just a fraction of a turn of the manipulator.
Tip: After creating a divot on the cell membrane, a slight tap on the manipulator also works to push the needle into the cell.
5. Before injecting, make sure the injector is set to inject 50 nL (see Nanoinject manual) and then press the inject button to dispense RNA into the oocyte. Oocytes may visibly inflate while pressing the inject button. We also use the foot peddle for injections to streamline injections.
6. We typically inject 10–20 cells. After injecting all cells, incubate the oocytes in ND-96 at 18 °C for 24–36 h.
Video 2. RNA injection of an oocyte. The video shows the various steps of RNA injection in an oocyte as outlined in section C.
D. TMRM labeling of oocytes
1. TMRM labeling is done on the day of the experimental recording. Labeled oocytes can be stored in the dark at 18 °C for the duration of the day. Storing for longer than 12 h will result in significant expression of unlabeled VSP, making the labeled protein unrepresentative of the whole pool of expressed protein.
2. Make the TMRM labeling mixture in a 2 mL round-bottom Eppendorf tube, containing 200 μL of ND-96 without gentamicin or pyruvate [ND-96(−)] plus 0.2 μL of the 25 mM TMRM (final concentration 25 μM). Vortex this mixture for 10 s.
a. Make the 25 mM TMRM stock with dry DMF and keep at -20 °C in the dark and in a container filled with DrieriteTM.
3. Transfer the oocytes to the TMRM labeling mixture with minimal liquid transfer to avoid dilution of the mixture.
Tip: Let gravity do the work and let the oocytes fall into the labeling solution instead of squirting them in.
4. Leave the oocytes to label in the dark and on ice for 20 min.
5. After labeling, wash the oocytes with 1 mL of ND-96(−). Repeat this wash five times.
6. Store the labeled oocytes in ND-96(−) in the dark and in an incubator at 18 °C until the experiment.
Note: While the oocytes are experimentally measured at room temperature, leaving the cells at room temperature instead of 18 °C will increase the amount of unlabeled protein on the membrane since protein expression does not stop and can lead to a deterioration of the quality of the oocytes. Therefore, labeled oocytes are left at 18 °C and are individually removed when ready to start an experiment.
E. Two-electrode voltage-clamp electrophysiology and fluorometry measurement of VSD motion
1. Set up the two-electrode voltage clamp rig.
a. Confirm that the Digidata, computer, amplifier, and LED are powered on. Confirm that the amplifier is in TEV (V1, Vi) mode and set to external command so the computer can control the amplifier. Before powering on the PMTs, make sure that they are shielded from light and the voltage is off.
b. Fill the bath chamber with the experimental measurement solution ND-96(−).
c. Fill the back two wells with 1 M NaCl. Add the silver-chloride wires from the bath/guard headstage to the wells.
The silver-chloride wires are connected to the P1 and P2 headstages with the V2 headstage wire spliced into the wire for the P1 headstage (Figure 7A, white and yellow wire, respectively).
Figure 7. VCF setup. A. An oocyte under voltage clamp showing the bath and headstage setup. The black arrows point to the wells containing the silver-chloride wires and agarose bridges. The yellow wire connects to the P1 and V2 headstages, and the white wire connects to the P2 headstage. B. Silver-chloride wires. Top: properly chlorinated; bottom: needs to be chlorinated as shiny metal patches are showing. C. Oocyte in the bath chamber with glass microelectrodes inserted; the surrounding gray mesh prevents the oocyte from rolling. D. Front of Dagan CA-1B amplifier in the TEV setting. The digital monitor is set to V2 showing the bath clamp; note that the amplifier internally inverts the command (i.e., a -80 mV holding shows +80 mV). To the right, the CLAMP is in the ON position and BATH/GUARD switch to ACTIVE.
d. Place the two glass agarose bridges into the bath to connect the bath chamber to each of the two back wells.
i. To make the agarose bridges, take the 3.5” glass capillaries and cut to approximately 1.5’’ length with a file or glass cutter (Figure 8A and B). Using a Bunsen burner and tweezers, flame the end of the glass so a bend will appear approximately 0.25” from the end. Cut longer pieces if you want taller bridges. Let it cool down. Using the tweezers, grab the bent end and flame the other side to bend that end about the same height. Keep the glass such that the bend happens in the same direction as the first bend. The end result should look like a bridge.
ii. Thread a piece of palladium wire through the open tube. Cut at a length such that the wire does not stick out from the glass. Tuck the wire into the bridge so neither end sticks out (Figure 8C and D).
iii. To prepare the agarose, measure enough agarose for a 2% solution in bridge buffer. We tend to make about 50 mL at a time. Transfer the solution to a low-sided beaker. The low side will make the transfer of the solution into the glass bridge easier.
iv. Melt the agarose in the microwave making sure it does not boil over, as that will change the concentration of the agarose. Place the melted agarose on a hot plate to keep it hot and melted.
v. The 3.5” glass capillaries come with a suction tube that can be connected on one side to the bridge and the other side to a syringe (Figure 8E and F).
vi. Once both sides are sealed, fill up the agarose solution into the bridge. Because air bubbles will disrupt the connection, we keep filling into the suction tube. Remove from the melted agarose and leave for approximately 10–20 s, letting the agarose in the bridge solidify; then, remove from the seal (Figure 8G).
vii. We prepare several bridges with inserted wires before melting the agarose. Let them all solidify completely and inspect for any air bubbles inside or at the ends. If air bubbles are visible, discard. If none are visible, test the bridges with the next step of the protocol. When stored in bridge solution, bridges last for years.
Figure 8. Making of agarose bridges. A. 3.5” glass capillary marked at 1.5”. B. The glass capillary from A cut with bend points marked 0.25” from either end. C. Top: palladium wire cut the length of the glass capillary. Bottom: the glass capillary bent at the 0.25” mark. D. Palladium wire threaded through the bent glass capillary. E. Bent and palladium-filled glass capillary attached to the suction tube, which is attached to a syringe. F. Close view of the bent and palladium-filled glass capillary attached to the suction tube. G. The final bridge made of a bent glass capillary filled with the bridge agarose solution and a palladium wire.
e. On the amplifier, turn the BATH/GUARD switch to ACTIVE (Figure 7D). If the bridges are good (both are needed), then the middle display under “Clamp” on the amplifier will read the number of nA needed to zero the voltage from the bath. If a single bridge is removed while the BATH/GUARD switch is ACTIVE, then the display will max out because of the open circuit and display a -1 or a 1.
f. Inspect the microelectrode holders for cleanliness and chlorination of the silver wire as verified by a dull gray appearance. Place the microelectrode holders onto the V1 and Vi headstages.
If part of the wire is shiny, remove the wire and soak in undiluted household bleach for a duration from 1 h to overnight. Be sure to rinse off excess bleach with diH2O before replacing it onto the electrode holder (Figure 7B).
g. Pull the glass microelectrodes using the Narishige puller on the appropriate settings that will give a resistance measure between 0.2 and 1 MΩ (see step E1n).
Tip: We only pull two glass microelectrodes before turning the puller off to allow it to cool for 15 min before pulling another two glass microelectrodes (if needed). This pause period helps maintain the quality of the microelectrode pulled by not overheating the filament.
Tip: Glass microelectrodes are fragile and should be handled with care. An easy microelectrode holder can be made from a 150 mm Petri dish with modeling clay pressed into the bottom to hold the electrodes (Figure 9).
Figure 9. Dish for holding glass microelectrodes. The holder is a 150 mm Petri dish with modeling clay pressed into the bottom.
h. Use a blunted 25-gauge needle to backfill the glass electrodes with 3 M KCl about halfway up the glass. With the sharp tip of the glass electrode down, flick out any bubbles that may have formed in the tip of the glass electrode, so that the air bubbles rise through the solution. The bubbles may be visible as they rise. Bubbles between the tip of the glass electrode and the wire will increase resistance leading to a poor clamp of the cell.
i. Securely fit the 3 M KCl-filled glass microelectrodes into their holders and place into the bath solution.
j. Open the Clampex software and set the holding voltage to -80 mV.
k. Lower both electrodes so that the tips are in the recording solution.
l. On the amplifier, press the V1 monitoring button. Adjust the V1 OFFSET knob until the electrode digital meter reads the applied holding voltage in the opposite sign (i.e., holding voltage is -80 mV, therefore the digital meter will read +80 mV).
m. Press the Vi monitoring button. Adjust the Vi OFFSET knob until the electrode digital meter reads the applied holding voltage in the opposite sign.
n. Press the V1 monitoring button and turn the “Z TEST” button in the V1 column to the ON position. The displayed value is the resistance of the V1 microelectrode. Press the Vi monitoring button and turn the “Z TEST” button in the Vi column to the ON position. The displayed value is the resistance of the Vi microelectrode.
i. Each microelectrode resistance should measure between 0.2 and 1 MΩ.
ii. If a microelectrode measures out of the resistance range, replace the glass microelectrode. A resistance higher than 1 MΩ could indicate air bubbles. Carefully remove the glass electrode and flick again to remove any remaining bubbles to lower the resistance (see step E1g).
o. To prevent light contamination of the fluorometry, turn off the lights in the room and turn on a desk lamp facing away from the VCF setup.
Tip: If unable to turn the room lights off, cloth blackout covers can be made to sit on top of the Faraday cage to protect the samples from the light.
2. After setting up the VCF rig, pull the electrodes out of the recording solution and then place the oocyte into the bath chamber. Gently roll the oocyte until the dark animal pole faces down onto the glass coverslip. We use a mesh to keep the oocytes from rolling around on the coverslip.
a. Roll by either pipetting up and down or through gentle pushing with an empty pulled glass microelectrode that has been flamed to create a closed blunted tip, like a hockey stick.
3. Bring the objective into focus on the oocyte. Because the oocyte will cover most of the field of view when using a 20× objective, aim to focus on the granules that give the dark pole its color since these will represent the bottom of the oocyte.
4. On the amplifier, press the monitoring button for V1-V2. If the offsets were properly adjusted as in step E1i–k, then the digital meter will read 0 volts, as both V1 and V2 are set to reading the bath clamp at 80 mV. Impale the V1 microelectrode into the oocyte membrane with as shallow a penetration as possible. We use the same technique here as we do when injecting the cells. The value of V1-V2 will increase as soon as the oocyte is impaled; this value is the oocyte resting membrane potential.
a. While the resting membrane potential is dependent on the health of the oocyte, the protein expressed, and the solution conditions of the oocyte, for our given experiments with Ci-VSP G214C in ND-96, the resting membrane potential should range from -20 to -60 mV. Values close to 0 mV indicate that the oocyte quality is poor. These poor-quality cells are discarded as they typically will not reliably hold a voltage clamp.
5. Repeat step E4, this time impaling the oocyte with the Vi microelectrode and using the V1-Vi digital meter (Figure 7C). Before the oocyte is impaled with Vi, both the monitors for V1-V2 and V1-Vi should read the resting membrane potential, as both V2 and Vi are still reading the bath clamp at 80 mV. As soon as the Vi microelectrode impales the oocyte, the V1-Vi digital meter will read 0 mV, as both V1 and Vi are reading the same oocyte internal values.
Tip: Simultaneous impalement makes it less likely that the oocyte will roll away. Create a divot with both electrodes and then tap the manual manipulators. That degree of force is usually enough to pop the electrode into the cell without damaging the cell.
6. Do a final focus of the objective on the oocyte (should just be small fine adjustments at this point).
7. Switch the microscope prism to send the emission light to the PMT port instead of the eye port. If using a blackout cover, lower the cover over the opening of the Faraday cage now.
8. On the amplifier, switch the CLAMP to the ON position (Figure 7D). If the oocyte is properly clamped, the V1 monitor should read 0 mV, as V1 monitors the inside of the cell held at a virtual ground, while the bath is voltage-clamped to the set holding voltage (-80 mV) as monitored by V2. Therefore, when the clamp is on, the V1-V2 monitor should read the oocyte holding voltage of -80 mV.
9. Turn the CLAMP GAIN knob to the ½ position. To confirm the integrity of the clamp and determine the clamp gain settings, run a single voltage pulse going from -80 mV to +150 mV for 125 ms and then back to -80 mV with the LED trigger off.
a. The Im (membrane current) signal should have minimal oscillations, and the waveform of the applied voltage pulse should be a tight square. If the Im signal shows oscillations, turn down the gain. If the voltage pulse shows a slow rise to the applied voltage step, turn the gain up. If adjusting the gain does not help, then the integrity of the oocyte membrane is likely not ideal, and another oocyte should be used.
10. Trigger the LED to turn on for 250 ms to establish the PMT baseline. Initially, the output signal from the PMT is close to 0 as there is no voltage in the system. Turn up the voltage on the PMT until the output signal measures around 6 V. This level provides a good signal while avoiding the potential saturation of the PMT (10 V).
11. Then, run a single pulse protocol, holding from -80 to +150 mV for 300 ms and then back to -80 mV with the LED set to 70% of max intensity triggered during the entire protocol (before, during, and after the voltage pulse).
a. If Ci-VSP G214C is expressed and labeled properly, a fluorescence decrease will be seen, corresponding to the +150 mV step as in Figure 5 of reference [6].
12. The stepwise VCF voltage protocol used in our lab consists of 10 mV steps starting at -150 mV and ending at 200 mV with a holding potential of -80 mV. The LED should be triggered at the same time for each step in the protocol.
Data analysis
1. The ΔF/F is calculated by measuring the amplitude of fluorescence change of each pulse and dividing by the initial value.
a. Start by opening the experimental pulse recordings in Clampfit.
b. Cursor 1 & Cursor 2 are locked with 100 ms between them (right click, Lock to Partner) and are moved to the fluorescence baseline before the voltage change is applied.
c. Cursor 3 & Cursor 4 are locked with 10 ms between them (right click, Lock to Partner) and are placed at the point where the fluorescence signal reaches the plateau before the end of the voltage step.
d. Under the Analyze tab, press the Statistics button (or Alt+S). This writes the mean value of each cursor pair into the results window.
e. Copy the values from the results window into Excel and calculate ΔF/F by ΔF/F = [(Fx - F0)/F0]. F0 is the initial fluorescence baseline value (Cursor 1–2) and Fx is the plateau of the fluorescence signal (Cursor 3–4).
2. To display the fluorescence signal, it is normalized in Clampfit by zeroing the fluorescence baseline.
a. Save this file as a .atf file (making sure that all signals and the whole trace options are selected to be saved).
b. Open this file in Igor: Data, Load Waves, Load Waves… In the Load Waves window that pops up, select General Text and select the file under the Path option.
c. Under Windows, select New Graph. In the New Graph popup window, the Y Wave selected is the fluorescence trace signal and the X Wave selected is the time signal.
3. In Igor, the normalized fluorescence signal is used to generate the kinetic fits. Both the activation and repolarization kinetics are obtained from either a single or double exponential.
a. To fit the activation kinetics:
i. The normalized fluorescence signal from step 2 is zoomed into the start of the fluorescence change.
ii. Cursor A is placed at the first decrease in the fluorescence signal.
iii. Cursor B is placed at the plateau of the fluorescence change.
iv. Under the Analysis tab, click Curve Fitting… In the popup window, under the Function and Data tab, select either the single exponential (exp) or the double exponential (dblexp).
v. Under the Data Options tab, select the Range as Cursors.
vi. Press Do It to run the fit.
b. To fit the repolarization kinetics, the steps are the same as in step 3a. However, move Cursor A to the very start of the fluorescence signal repolarization where the voltage pulse has returned to the -80 mV holding potential. And move Cursor B to the point where the fluorescence signal has returned to baseline.
c. To determine whether a single or double exponential fit is more appropriate, start with a single exponential and visually determine whether the fit follows the data. If parts of the data are not properly represented by the fit, try a double exponential. In general, a double exponential is almost always better than a single, but to justify the additional parameters, a significant improvement of the fit is needed. See Figure 2d in [5] for an example. To quantitate whether the improvement is sufficient, we use residuals (data point minus the fit).
4. F/V curves are created by taking the amplitude of fluorescence change of the normalized fluorescence signal and plotting that value against the corresponding voltage value.
a. The fluorescence trace is opened in Clampfit. Cursors 1 and 2 are placed to select the last 10 ms of the voltage pulse. Cursor 3 is placed toward the end of the voltage pulse.
i. Press the I/V button.
ii. In the resulting I-V window popup, set cursor 3 as the X-axis so the values correspond to the actual applied voltage.
iii. The Y-axis values correspond to the mean of the value between cursor 1 and 2 in the fluorescence signal.
iv. After pressing ok, this writes the F/V values into the results window.
b. The F/V values are then imported into Igor to be normalized from -1 to 0 by dividing all the fluorescence signal values by the fluorescence signal value at the +200 mV stimulus.
c. The F/V curves are then fit with either a single or double Boltzmann sigmoid equation (Igor lists this option just as a sigmoid) to obtain the V1/2 and slope values.
i. The double sigmoid equation is manually added into Igor using the following equation:
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m
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x
1
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e
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x
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a
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ii. To overlay multiple F/Vs, we recommend re-normalizing the curves to the individual curve fits. This will give exact -1 and 0 values at the ends of the curves, facilitating the visual observation and comparison of the V1/2 and slope values.
Validation of protocol
This protocol or parts of it was used in the following research article:
• Rayaprolu et al. [6]. Hydrophobic residues in S1 modulate enzymatic function and voltage sensing in voltage-sensing phosphatase. Journal of General Physiology. (Figure 5, Figure S2, and Figure S3)
General notes and troubleshooting
General notes
1. The voltage clamping procedures outlined in this protocol are specific to the Dagan CA-1B amplifier, which is no longer manufactured. There are other models of voltage clamp amplifiers on the market, such as the currently available Axoclamp 900A Amplifier (Molecular Devices), TEC-03X, and TEC-10CX (Npi Electronic). While the principles of two-electrode voltage clamp are the same, the specifics of user interface change. Please follow the directions for setting up two-electrode voltage clamping per each specific manufacturer.
2. This protocol does not cover the setting up of a solution exchange system for the bath chamber as it was not needed in the experiments detailed here. However, the rapid changing of the bath solution surrounding the oocyte is a route experimental variable that can easily be set up in any VCF system.
3. This protocol details fluorometry using a cysteine crosslinking fluorophore (TMRM). Other fluorescent reporters are also suitable for VCF including Alexa dyes, Monobromo(trimethylammonio)bimane bromide (qBBr), or fluorescein. Additionally, to probe sites not extracellularly available, ANAP [3-(6-acetylnaphthalen-2-ylamino)-2-aminoprop anoic acid] [14], a fluorescent unnatural amino acid incorporated by genetic means, can be used.
4. VSP does not contain any extracellularly available native cysteines; however, other proteins of interest may have extracellularly available native cysteines that may also be labeled with TMRM. Two problems may arise from this native labeling. The labeled native cysteine may respond to voltage and generate its own VCF signal. If this signal is not wanted, we recommend mutating the native cysteine residues to create a cysteine-less background. The cysteine-less protein must be tested to measure the extent to which cysteine replacement did or did not affect function. If the native cysteine labels but does not change fluorescence with voltage, then the experimenter needs to decide if the background fluorescence is low enough for a true voltage-dependent signal to be detected. If so, then leave the native cysteine. If the background fluorescence is too high to see signals, we again recommend generating a cysteine-less protein. If a cysteine-less protein is not an option because it is unacceptably functionally changed, we recommend testing other attachment chemistries like methanethiosulfonate (MTS) or other fluorophores, since the size of the fluorophore will impact the labeling efficiency depending on the exposure of the cysteine. Another option is to change the method of the fluorophore incorporation to a genetic incorporation of an unnatural amino acid such as ANAP [14].
Troubleshooting
Problem 1: Poor oocyte health.
Possible cause: Improper incubating solution or issues with overexpression of the protein.
Solution: Ensure that the ND-96 solution is not contaminated and is at the correct pH and osmolarity. If the ND-96 solution is good, then lowering the incubation temperature to slow the trafficking of the protein can be useful. In addition, injecting less RNA and empirically determining incubation times is useful. While we use the cells for a week after processing, their health can decline during those 7 days. Using cells the day after processing will usually result in better experiments.
Problem 2: Poor protein expression, small or no fluorescence change.
Possible cause: Poor mRNA quality or issues with the protein expression.
Solution: Make sure that the mRNA is produced under RNase-free conditions to prevent degradation. Check for degradation using RNA gels. The concentration of the solution will not necessarily reflect degradation. Store RNA at ≤-20 °C and keep on ice while using. Make sure the DNA linearization reaction is complete. The RNA polymerase is very processive and will not easily fall off any contaminating plasmid, therefore producing nonfunctional RNA. Increasing incubation time and temperature may also improve protein expression.
Problem 3: Pink hue on oocytes after labeling.
Possible cause: The oocytes have been over labeled and the TMRM in the membrane creates a high background of fluorescence.
Solution: Take care to label the oocytes on ice for no longer than 20 min. Increase wash time and volume. TMRM will also label the membranes when they are compromised. If the proper labeling conditions have been followed and the pink hue is still present, the plasma membranes are absorbing the dye and are unlikely to clamp properly.
Acknowledgments
This protocol is related to the following paper: Rayaprolu et al. Hydrophobic residues in S1 modulate enzymatic function and voltage sensing in voltage-sensing phosphatase https://doi.org/10.1085/jgp.202313467. We thank Y. Okamura for providing the Ci-VSP cDNA. This work was supported by funds from the National Institute of General Medical Science of the National Institutes of Health grants R01GM111685 (S.C.K.) and National Science Foundation grant 2310489 (S.C.K.).
Competing interests
Authors declare no competing financial interests.
Ethical considerations
All animal tissue used was purchased from approved suppliers of Xenopus laevis oocytes.
References
Mannuzzu, L. M., Moronne, M. M., and Isacoff, E.Y. (1996). Direct physical measure of conformational rearrangement underlying potassium channel gating. Science. 271(5246): 213–216.
Cha, A. and Bezanilla, F. (1997). Characterizing voltage-dependent conformational changes in the Shaker K+ channel with fluorescence. Neuron. 19(5): 1127–1140.
Dürr, K. L., Abe, K., Tavraz, N. N. and Friedrich, T. (2009). E2P State Stabilization by the N-terminal Tail of the H,K-ATPase β-Subunit Is Critical for Efficient Proton Pumping under in Vivo Conditions. J Biol Chem. 284(30): 20147–20154.
Young, V. C. and Artigas, P. (2021). Displacement of the Na+/K+ pump’s transmembrane domains demonstrates conserved conformational changes in P-type 2 ATPases. Proc Natl Acad Sci USA. 118(8): e2019317118.
Kohout, S. C., Ulbrich, M. H., Bell, S. C. and Isacoff, E. Y. (2008). Subunit organization and functional transitions in Ci-VSP. Nat Struct Mol Biol. 15(1): 106–108.
Rayaprolu, V., Miettinen, H. M., Baker, W. D., Young, V. C., Fisher, M., Mueller, G., Rankin, W. O., Kelley, J. T., Ratzan, W. J., Leong, L. M., et al. (2024). Hydrophobic residues in S1 modulate enzymatic function and voltage sensing in voltage-sensing phosphatase. J Gen Physiol. 156(7): e202313467.
Murata, Y., Iwasaki, H., Sasaki, M., Inaba, K. and Okamura, Y. (2005). Phosphoinositide phosphatase activity coupled to an intrinsic voltage sensor. Nature. 435(7046): 1239–1243.
Villalba-Galea, C. A., Sandtner, W., Starace, D. M. and Bezanilla, F. (2008). S4-based voltage sensors have three major conformations. Proc Natl Acad Sci USA. 105(46): 17600–17607.
Villalba-Galea, C. A., Miceli, F. and Bezanilla, F. (2009). Uncoupling Of The Phosphatase Produces A Deeper Relaxation Of Ci-VSP. Biophys J. 96(3): 370a.
Kohout, S. C., Bell, S. C., Liu, L., Xu, Q., Minor, D. L. and Isacoff, E. Y. (2010). Electrochemical coupling in the voltage-dependent phosphatase Ci-VSP. Nat Chem Biol. 6(5): 369–375.
Mizutani, N., Kawanabe, A., Jinno, Y., Narita, H., Yonezawa, T., Nakagawa, A. and Okamura, Y. (2022). Interaction between S4 and the phosphatase domain mediates electrochemical coupling in voltage-sensing phosphatase (VSP). Proc Natl Acad Sci USA. 119(26): e2200364119.
Castle, P. M., Zolman, K. D. and Kohout, S. C. (2015). Voltage-sensing phosphatase modulation by a C2 domain. Front Pharmacol. 6: e00063.
Paixao, I. C., Mizutani, N., Matsuda, M., Andriani, R. T., Kawai, T., Nakagawa, A., Okochi, Y. and Okamura, Y. (2023). Role of K364 next to the active site cysteine in voltage-dependent phosphatase activity of Ci-VSP. Biophys J. 122(11): 2267–2284.
Kalstrup, T. and Blunck, R. (2013). Dynamics of internal pore opening in KV channels probed by a fluorescent unnatural amino acid. Proc Natl Acad Sci USA. 110(20): 8272–8277.
Article Information
Publication history
Received: Nov 11, 2024
Accepted: Dec 30, 2024
Available online: Jan 19, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Biochemistry > Protein > Fluorescence
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5,213 | https://bio-protocol.org/en/bpdetail?id=5213&type=0 | # Bio-Protocol Content
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This is an In Press version of the protocol that has not yet been assigned to an issue.
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HPLC Analysis of tRNA‐Derived Nucleosides
XC Xingxing Chen
FX Fu Xu
In Press, Available online: Jan 19, 2025
DOI: 10.21769/BioProtoc.5213 Views: 18
Reviewed by: Ritu GuptaJaveena Hussain Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in RNA Dec 2024
Abstract
Transfer RNAs (tRNAs), the essential adapter molecules in protein translation, undergo various post-transcriptional modifications. These modifications play critical roles in regulating tRNA folding, stability, and codon–anticodon interactions, depending on the modified position. Methods for detecting modified nucleosides in tRNAs include isotopic labeling combined with chromatography, antibody-based techniques, mass spectrometry, and high-throughput sequencing. Among these, high-performance liquid chromatography (HPLC) has been a cornerstone technique for analyzing modified nucleosides for decades. In this protocol, we provide a detailed, streamlined approach to purify and digest tRNAs from yeast cells and analyze the resulting nucleosides using HPLC. By assessing UV absorbance spectra and retention times, modified nucleosides can be reliably quantified with high accuracy. This method offers a simple, fast, and accessible alternative for studying tRNA modifications, especially when advanced technologies are unavailable.
Key features
• A streamlined protocol for purifying total tRNAs from yeast cells.
• Adaptable for other RNA species and organisms, provided sufficient input material.
• Enables the quantification of approximately 20 types of tRNA modifications.
• Offers a cost-effective and rapid alternative for analyzing tRNA modifications by HPLC method.
Keywords: tRNA modifications HPLC Retention time UV absorbance Epitranscriptome Yeast
Background
The landscape of RNA modifications across the transcriptome, known as the epitranscriptome, is receiving increasing attention. To date, over 170 distinct types of RNA modifications have been identified [1]. These modifications play pivotal roles in regulating all aspects of RNA biology, from transcription and processing to degradation, ultimately influencing cell fate. Among all RNA species, tRNA stands out as one of the most extensively modified, with an average of 13 modifications per molecule [2]. These modifications encompass a wide range of chemical alterations, from simple methylations to complex structures like pseudouridylation. Deficiencies in tRNA modifications or the enzymes responsible for them have been linked to various human diseases, including neurological disorders and cancer [3]. Thus, studying tRNA modifications is critical for advancing our understanding of human health.
Various methods are available for the quantitative analysis of tRNA modifications. Among these, mass spectrometry, regarded as the gold standard for tRNA modification identification, is widely used in the field. Additionally, emerging high-throughput sequencing technologies, such as mim-tRNAseq and nano-tRNAseq, are revolutionizing the study of tRNA modifications by enabling single-nucleoside resolution analysis at the epitranscriptome level [4,5]. Among the classic methods, for decades HPLC has been a cornerstone technique for analyzing modified nucleosides, remaining a valuable tool in RNA modification research. Here, we present a modified version of the protocol originally described by Gehrke et al. [6]. This updated protocol employs a C30 reverse-phase column coated with a 30-carbon-long alkyl chain, offering a more hydrophobic separation phase compared to the previously used C18 column, which features an 18-carbon-long alkyl chain. Additionally, we provide a streamlined protocol for purifying tRNAs from yeast cells using diethylaminoethyl cellulose (DE52) for subsequent HPLC analysis. The cellulose matrix exhibits weak ion exchange properties, enabling the separation of RNAs of different sizes under varying salt concentrations [7]. With this approach, we can reliably separate and identify more than 20 types of modified nucleosides with high accuracy and confidence.
Materials and reagents
Biological materials
1. Saccharomyces cerevisiae
Reagents
1. Synthetic complete yeast growth medium (Formedium, catalog number: CSC0201)
2. Nuclease-free water (Invitrogen, catalog number: 10977-035)
3. NaCl (Sigma-Aldrich, catalog number: S9888)
4. Water-saturated phenol (AppliChem, catalog number: A1624)
5. Chloroform (Sigma-Aldrich, catalog number: 650498)
6. Ethanol (Sigma-Aldrich, catalog number: 1009861000)
7. 1 M Tris-HCl, pH 7.4 (Santa Cruz, catalog number: SC-301950)
8. 1 M Tris-HCl, pH 8.3 (Fisher Scientific, catalog number: T1083)
9. Qubit RNA Broad Range kit (Thermo Fisher, catalog number: Q10210)
10. Isopropanol (Sigma-Aldrich, catalog number: 1096341011)
11. DMSO (Sigma-Aldrich, catalog number: 34869)
12. Diethylaminoethyl cellulose (DE52) (Biophoretics, catalog number: B45059)
13. Nuclease P1 (Sigma-Aldrich, catalog number: N8630)
14. Bacterial alkaline phosphatase (BAP) (Sigma-Aldrich, catalog number: P4252)
15. ZnCl2 (Sigma-Aldrich, catalog number: 39059)
16. 1 M sodium acetate, pH 5.0 (Thermo Fisher, catalog number: J60964.AK)
17. NH4H2PO4 solution (Sigma-Aldrich, catalog number: 216003)
18. NH4OH solution (Sigma-Aldrich, catalog number: 221228)
19. H3PO4 solution (Sigma-Aldrich, catalog number: 695017)
20. Methanol (Honeywell Chemicals, catalog number: 34966)
21. Acetonitrile (Honeywell Chemicals, catalog number: 34881)
Solutions
1. 0.9% NaCl (w/v)
2. 10 mM ZnCl2
3. DE52 binding buffer (see Recipes)
4. DE52 solution (see Recipes)
5. tRNA elution buffer (see Recipes)
6. Nuclease P1 solution (see Recipes)
7. HPLC solvent A (see Recipes)
8. HPLC solvent B (see Recipes)
9. HPLC solvent C (see Recipes)
Recipes
1. 0.9% NaCl
Store at RT.
Reagent Final concentration Amount
NaCl 0.9% (w/v) 9 g
Nuclease-free H2O n/a 1,000 mL
2. 10 mM ZnCl2
Store at -20 °C. Stable for at least five years.
Reagent Final concentration Amount
0.1 M ZnCl2 0.01 M 100 μL
Nuclease-free H2O n/a 900 μL
3. DE52 binding buffer
Store at RT. Stable for at least one year.
Reagent Final concentration Amount
1 M Tris-HCl, pH 7.4 0.1 M 100 mL
NaCl 0.1 M 5.844 g
Nuclease-free H2O n/a 900 mL
4. DE52 solution
Store at 4 °C. Stable for at least six months.
Reagent Final concentration Amount
DE52 0.33 g/mL (w/v) 100 g
DE52 binding buffer n/a 300 mL
5. tRNA elution buffer
Store at RT. Stable for at least one year.
Reagent Final concentration Amount
1 M Tris-HCl, pH 7.4 0.1 M 100 mL
NaCl 1 M 58.44 g
Nuclease-free H2O n/a 900 mL
6. Nuclease P1 solution
Store at -20 °C. Stable for at least one year.
Reagent Final concentration Amount
Nuclease P1 >250 units /1 mL 1 vial (>250 units)
1 M sodium acetate, pH 5.0 30 mM 30 μL
Nuclease-free H2O n/a 970 μL
7. HPLC solvent A
Prepare before use. Adjust pH to 5.3 with NH4OH and H3PO4.
Reagent Final concentration Amount
1 M NH4H2PO4 0.01 M 10 mL
Methanol 2.5% (v/v) 25 mL
Milli-Q H2O n/a 965 mL
8. HPLC solvent B
Prepare before use. Adjust pH to 5.1 with NH4OH and H3PO4.
Reagent Final concentration Amount
1 M NH4H2PO4 0.01 M 10 mL
Methanol 20% (v/v) 200 mL
Milli-Q H2O n/a 790 mL
9. HPLC solvent C
Prepare before use.
Reagent Final concentration Amount
1 M NH4H2PO4 0.01 M 10 mL
Acetonitrile 35% (v/v) 350 mL
Milli-Q H2O n/a 640 mL
Laboratory supplies
1. Disposable gloves
2. Milli-Q water
3. 15 mL centrifuge tubes (Corning, catalog number: 430052)
4. 50 mL centrifuge tubes (Corning, catalog number: 352070)
5. 1.5 mL micro-centrifuge tubes (Eppendorf, catalog number: 0030120086)
6. 1 L glass bottles (Thermo Fisher, catalog number: FB8001000)
7. HPLC sample tube (Waters, catalog number: 186000273)
8. HPLC sample tube cap (Waters, catalog number: 186000305)
9. HPLC sample insert (Waters, catalog number: WAT094170)
10. Polypropylene column (Bio-Rad, catalog number: 731-1550)
11. Reversed phase aqueous C30 column (Phenomenex, catalog number: CH0-5690)
Equipment
1. Single channel pipette set (Sigma-Aldrich, catalog number: EP3123000918-1EA)
2. Qubit fluorometer (Thermo Fisher, catalog number: Q33238; or similar)
3. 30 °C shaking incubator (INFORS-HT, model: Ecotron; or similar)
4. Cell density photometer (Implen, model: DiluPhotometer; or similar)
5. Centrifuge for 50 mL conical tube (Heraeus, model: Multifuge X3R; or similar)
6. Benchtop centrifuge for 1.5 mL centrifuge tube (Eppendorf, model: 5427R; or similar)
7. Benchtop mixer (Heidolph Instrument, model: 545-10000-00; or similar)
8. pH meter (Mettler Toledo, model: 30266626; or similar)
9. HPLC separation module (Waters, model: 2695)
10. PDA detector (Waters, model: 2996)
Software and datasets
1. Empower 3 FR4 (Waters, Version 3 Feature Release 4, released in 2017); requires a license
Procedure
A. tRNA isolation
1. Harvest 100 units (100 mL of yeast culture at OD600 = 1) of yeast cells grown in synthetic complete yeast growth medium at log phase in 50 mL centrifuge tubes. Centrifuge the tubes at 1,000× g for 3 min at room temperature (RT; 25 °C) and remove the supernatant.
2. Wash the cells once with 3 mL of 0.9% NaCl. Centrifuge at 1,000× g at RT for 3 min and remove the supernatant.
3. Resuspend the cells in 8 mL of phenol and vortex at 1,200 rpm using a benchtop mixer for 30 min at RT.
4. Add 400 μL of chloroform and vortex at 1,200 rpm for an additional 15 min at RT. Centrifuge at 12,000× g for 20 min at 4 °C.
5. Collect the water phase in a 50 mL conical tube, add 4 mL of water-saturated phenol, and vortex at 1,200 rpm for 15 min at RT.
6. Centrifuge at 12,000× g for 20 min at 4 °C. Collect the water phase and mix it with 2.5 volumes of ethanol by inverting the tubes three times.
7. Precipitate at -80 °C for 3 h or -20 °C overnight.
8. Centrifuge at 12,000× g for 20 min at 4 °C. Remove the supernatant and dissolve the RNA pellet in 5 mL of DE52 binding buffer.
9. Prepare a DE52 cellulose column by adding 5 mL of thoroughly mixed DE52 solution to a polypropylene column. Let the liquid pass through the column for approximately 5 min by gravity at RT.
10. Add the resuspended RNA pellet to the DE52 cellulose column and let the liquid pass through the column by gravity at RT.
11. Add 7 mL of DE52 binding buffer to the column and wait 5 min. Repeat the step once more.
12. Elute the RNA by adding 7 mL of tRNA elution buffer and collect the elute in a 15 mL centrifuge tube. Elute the tRNA once more by adding the elute back to the column to increase the yield.
13. Add 5 mL of isopropanol to the elute and mix by inverting the tube three times. Precipitate tRNA at -20 °C for at least 3 h.
14. Centrifuge at 12,000× g for 20 min at 4 °C to pellet tRNA. Discard the supernatant and resuspend the pellet in 1 mL of 70% EtOH.
15. Transfer the resuspension to a 1.5 mL centrifuge tube and spin at 16,000× g for 5 min at 4 °C to pellet tRNA.
16. Dissolve the tRNA in 50 μL of nuclease-free water. Measure the tRNA concentration with Qubit RNA Broad Range kit. Approximately 200 μg of tRNA can be obtained from 100 units of yeast cells.
17. The tRNA can be stored at -80 °C for up to one week or continued with tRNA digestion.
B. tRNA digestion
1. Perform nuclease P1 digestion by preparing the following mix and incubating at 37 °C for 16 h.
50 μg tRNA x μL
10 mM ZnCl2 5 μL
Nuclease P1 solution 10 μL
Nuclease-free H2O Add to 50 μL
2. Dephosphorylate the P1-digested tRNA by preparing the following mix and incubating at 37 °C for 2 h. Samples are ready for HPLC analysis.
Digested tRNA 50 μL
0.5 M Tris-HCl pH 8.3 mM 20 μL
BAP 4 μL
C. HPLC analysis
1. Prepare 1 L of each HPLC solvent (A, B, and C; see Recipes).
2. Put the solvent tubes (marked A, B, or C) into the corresponding HPLC solvent. Solvent tube D will not be used and can be kept in water.
3. Turn on both the HPLC separation module and the PDA detector.
4. Wet prime the HPLC system with a flow rate of 4 mL/min for 4 min.
5. Turn on the degas function.
6. Open Empower 3 software. Set up the gradient according to Table 1.
Table 1. Gradient of the HPLC method.
Step Time (min) Flow (mL/min) %A %B %C %D Curve
1 1.00 100 0 0 0
2 12 1.00 100 0 0 0 11
3 20 1.00 90 10 0 0 6
4 25 1.00 75 25 0 0 6
5 32 1.00 40 60 0 0 6
6 36 1.00 38 62 0 0 6
7 45 1.00 0 100 0 0 6
8 80 1.00 0 0 100 0 6
9 95 1.00 100 0 0 0 6
10 175 0.05 34 33 33 0 6
7. Set up the auto-injection volume to 60 μL and the running time to 140 min for each sample.
8. Transfer the digested tRNAs to the 2 mL HPLC sample tube with the sample insert and cap.
9. Update the sample inject table starting with nuclease-free water as the first sample and DMSO as the second to clean the C30 column.
10. Continue running the digested tRNA samples on the HPLC.
Data analysis
This protocol is adapted from the original HPLC chromatography method [6]. Each nucleoside is characterized by a unique UV absorbance spectrum and retention time, which are used for its identification and quantification. The retention times of the nucleosides are shown in Figure 1A, while the UV absorbance spectra for ncm5U, mcm5U, mcm5s2U, and yW are displayed in Figure 1B. Additional UV absorbance spectra can be found in this chapter [6]. For HPLC analysis, we typically include tRNAs isolated from cells deficient in the formation of the target nucleosides as a positive control. For instance, tRNAs from elp3Δ cells, which are deficient in producing wobble uridine modifications, are used to analyze ncm5U, mcm5U, and mcm5s2U [8].
Figure 1. HPLC analysis of tRNA-derived nucleosides. A. Representative HPLC chromatography of modified nucleosides derived from yeast total tRNAs. B. Representative UV absorbance spectra of modified nucleosides. Abbreviations: absorbance unit (A.U.); 5-carbamoylmethyluridine (ncm5U); 5-methoxycarbonylmethyluridine (mcm5U); 5-methoxycarbonylmethyl-2-thiouridine (mcm5s2U); wybutosine (yW). Full names of the rest of the abbreviations can be found in [1].
Validation of protocol
This protocol has been used and validated in the following research articles:
• Xu et al. [8]. Sod1-deficient cells are impaired in formation of the modified nucleosides mcm5s2U and yW in tRNA. RNA (Figures 1–4, Supplementary Figure S1, and S2).
• Xu et al. [9]. SSD1 suppresses phenotypes induced by the lack of Elongator-dependent tRNA modifications. PLOS Genetics (Supplementary Figure 5).
• Xu et al. [10]. Identification of factors that promote biogenesis of tRNACGASer. RNA Biology (Supplementary Figure S2).
• Xu et al. [11]. Yeast Elongator protein Elp1p does not undergo proteolytic processing in exponentially growing cells. MicrobiologyOpen (Figure 4, panel B).
• Chen et al. [12]. Elongator complex influences telomeric gene silencing and DNA damage response by its role in wobble uridine tRNA modification. PLOS Genetics (Figure 5).
• Chen et al. [13]. Defects in tRNA modification associated with neurological and developmental dysfunctions in Caenorhabditis elegans elongator mutants. PLOS Genetics (Figure 2 and supplementary Figure S1–S3).
• Huang et al. [14]. A genome-wide screen identifies genes required for formation of the wobble nucleoside 5-methoxycarbonylmethyl-2-thiouridine in Saccharomyces cerevisiae. RNA (Figure 2).
• Björk et al. [15]. A conserved modified wobble nucleoside (mcm5s2U) in lysyl-tRNA is required for viability in yeast. RNA (Figure 5).
• Esberg et al. [16]. Elevated levels of two tRNA species bypass the requirement for elongator complex in transcription and exocytosis. Molecular Cell (Supplementary Table 1).
• Huang et al. [17]. An early step in wobble uridine tRNA modification requires the Elongator complex. RNA (Table 1 and Figure 3).
General notes and troubleshooting
1. Various media are available for growing yeast cells, including YEPD and synthetic dropout media with additional supplements. We observed lower background noise during HPLC analysis of tRNA nucleosides from cells grown in synthetic dropout medium.
2. Digested tRNA can be stored at -80 °C and analyzed later. No significant differences were observed in nucleoside analysis for samples stored at -80 °C for up to one week. For longer storage, the stability of modified nucleosides should be evaluated.
3. When a positive control strain is available, it is essential to include tRNAs isolated from the control strain. The retention time of each nucleoside may vary by seconds to minutes, depending on the precision of HPLC solvent preparation.
4. To maintain optimal separation of nucleosides, we typically replace the C30 column after analyzing approximately 50 samples or when the retention times deviate significantly from the expected values.
5. While nucleoside spectra are generally stable, we noticed increased background noise when the UV lamp in the PDA detector has been used for a long period.
6. We typically use 50 μg of tRNA as the starting material for analyzing wobble uridine modifications. Smaller input amounts may suffice for abundant modified nucleosides such as m1A, m5C, m1G, ac4C, and t6A.
7. The methanol concentration in HPLC solvent A can be increased to 6% (v/v) for the detection of m3C [10].
8. This protocol can also be applied to analyze tRNA nucleosides derived from other organisms, provided sufficient input material is available.
Acknowledgments
We thank Dr. Anders S. Byström and his laboratory members for general support. The protocol was used in articles listed in the section “Validation of protocol”.
Competing interests
The authors declare no competing interests.
References
Cappannini, A., Ray, A., Purta, E., Mukherjee, S., Boccaletto, P., Moafinejad, S. N., Lechner, A., Barchet, C., Klaholz, B. P., Stefaniak, F., et al. (2024). MODOMICS: a database of RNA modifications and related information. 2023 update. Nucleic Acids Res. 52: D239–D244.
Phizicky, E. M. and Hopper, A. K. (2023). The life and times of a tRNA. RNA. 29(7): 898–957.
Chujo, T. and Tomizawa, K. (2021). Human transfer RNA modopathies: diseases caused by aberrations in transfer RNA modifications. FEBS J. 288(24): 7096–7122.
Shaw, E. A., Thomas, N. K., Jones, J. D., Abu-Shumays, R. L., Vaaler, A. L., Akeson, M., Koutmou, K. S., Jain, M. and Garcia, D. M. (2024). Combining Nanopore direct RNA sequencing with genetics and mass spectrometry for analysis of T-loop base modifications across 42 yeast tRNA isoacceptors. Nucleic Acids Res. 52(19): 12074–12092.
Behrens, A., Rodschinka, G. and Nedialkova, D. D. (2021). High-resolution quantitative profiling of tRNA abundance and modification status in eukaryotes by mim-tRNAseq. Mol Cell. 81(8): 1802–1815 e1807.
Gehrke, C. W. and Kuo, K. C. (1990). Chapter 1 Ribonucleoside Analysis by Reversed-Phase High Performance Liquid Chromatography. In: Gehrke, C. W. and Kuo, K. C. T. (Eds.) J Chromatogr Library. Elsevier, A3–A71.
Kawade, Y., Okamoto, T. and Yamamoto, Y. (1963). Fractionation of soluble RNA by chromatography on DEAE ion exchangers. Biochem Biophys Res Commun. 10: 200–203.
Xu, F., Bystrom, A. S. and Johansson, M. J. O. (2024). Sod1-deficient cells are impaired in formation of the modified nucleosides mcm(5)s(2)U and yW in tRNA. RNA. 30(12): 1586–1595.
Xu, F., Bystrom, A. S. and Johansson, M. J. O. (2019). SSD1 suppresses phenotypes induced by the lack of Elongator-dependent tRNA modifications. PLoS Genet. 15(8): e1008117.
Xu, F., Zhou, Y., Bystrom, A. S. and Johansson, M. J. O. (2018). Identification of factors that promote biogenesis of tRNA(CGA)(Ser). RNA Biol. 15(10): 1286–1294.
Xu, H., Bygdell, J., Wingsle, G. and Bystrom, A. S. (2015). Yeast Elongator protein Elp1p does not undergo proteolytic processing in exponentially growing cells. Microbiologyopen. 4(6): 867–878.
Chen, C., Huang, B., Eliasson, M., Ryden, P. and Bystrom, A. S. (2011). Elongator complex influences telomeric gene silencing and DNA damage response by its role in wobble uridine tRNA modification. PLoS Genet. 7(9): e1002258.
Chen, C., Tuck, S. and Bystrom, A. S. (2009). Defects in tRNA modification associated with neurological and developmental dysfunctions in Caenorhabditis elegans elongator mutants. PLoS Genet. 5(7): e1000561.
Huang, B., Lu, J. and Bystrom, A. S. (2008). A genome-wide screen identifies genes required for formation of the wobble nucleoside 5-methoxycarbonylmethyl-2-thiouridine in Saccharomyces cerevisiae. RNA. 14(10): 2183–2194.
Bjork, G. R., Huang, B., Persson, O. P. and Bystrom, A. S. (2007). A conserved modified wobble nucleoside (mcm5s2U) in lysyl-tRNA is required for viability in yeast. RNA. 13(8): 1245–1255.
Esberg, A., Huang, B., Johansson, M. J. and Bystrom, A. S. (2006). Elevated levels of two tRNA species bypass the requirement for elongator complex in transcription and exocytosis. Mol Cell. 24(1): 139–148.
Huang, B., Johansson, M. J. and Bystrom, A. S. (2005). An early step in wobble uridine tRNA modification requires the Elongator complex. RNA. 11(4): 424–436.
Article Information
Publication history
Received: Nov 30, 2024
Accepted: Jan 5, 2025
Available online: Jan 19, 2025
Copyright
© 2025 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
Cancer Biology
Biological Sciences > Biological techniques
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5,214 | https://bio-protocol.org/en/bpdetail?id=5214&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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Peer-reviewed
A Novel Gene Stacking Method in Plant Transformation Utilizing Split Selectable Markers
GY Guoliang Yuan *
MI Md Torikul Islam *
GT Gerald A. Tuskan
XY Xiaohan Yang
(*contributed equally to this work)
In Press, Available online: Jan 16, 2025
DOI: 10.21769/BioProtoc.5214 Views: 48
Reviewed by: Xiaofei LiangHao Chen Anonymous reviewer(s)
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Cited by
Original Research Article:
The authors used this protocol in Communications Biology May 2023
Abstract
Gene stacking, the process of introducing multiple genes into a single plant to enhance desired traits, is essential for plant genetic improvement through both conventional breeding and genetic transformation. In general, transformation-based gene stacking can be achieved through either co-transformation to simultaneously introduce multiple genes or sequential multi-round transformation. While co-transformation is generally faster and more efficient than sequential multi-round transformation, it often requires two selectable marker genes, which confer resistance to antibiotics, for selecting transgenic events. However, in most cases, there is only one best selectable marker gene for a specific plant species or genotype. Also, it is harder to optimize the concentrations of two antibiotics for co-transformation than using one antibiotic for selecting transgenic events. To overcome this challenge, we recently developed an innovative split selectable marker system for plant co-transformation, allowing the use of one selectable marker gene to select transgenic events. This method involves constructing two binary vectors, each carrying a subset of genes of interest and a partial fragment of the selectable marker gene, which is connected to a partial intein fragment. Following Agrobacterium-mediated co-transformation, plants harboring both binary vectors are selected using a single antibiotic, such as kanamycin. This split-marker system can be used to co-transform multiple genes into both herbaceous and woody plants, accelerating genetic improvement of polygenic traits or integrative improvement of multiple traits to simultaneously increase crop yield and quality.
Key features
• Developed an intein-mediated split selectable marker system for efficient gene stacking in plants.
• Utilizes a single antibiotic for identifying transgenic events, simplifying the selection process of co-transformation compared to traditional methods.
• Applicable to both herbaceous and woody plant species for co-transforming multiple genes.
• Enhances scalability and feasibility of gene stacking in plant genetic engineering and crop improvement initiatives.
Keywords: Intein Selectable marker Protein splicing Plant transformation Arabidopsis Poplar
Graphical overview
Background
Gene stacking is a pivotal strategy in modern agriculture, involving the integration of multiple beneficial genes into a single organism to simultaneously enhance traits such as yield, disease resistance, stress tolerance, and nutritional content [1]. This approach is particularly valuable in the metabolic engineering of plants due to the intricate nature of metabolic processes and biochemical pathways, which typically involve complex interactions among multiple genes [2]. Gene stacking in plants can be achieved through several methods. First, hybrid stacking involves cross-hybridizing a plant containing one or more transgenes with another plant carrying different transgenes, resulting in the development of a multi-stack hybrid through iterative hybridization. Second, co-transformation entails transforming a plant with two or more independent transgenes, each in separate DNA constructs delivered simultaneously to the plant. Third, sequential multi-round transformation involves re-transformation of a plant already harboring a transgene with additional transgenes [3].
Co-transformation has proven to be a highly promising approach for introducing multiple genes into plants [4]. Typically, co-transformation involves using two different selectable marker genes simultaneously, which must be compatible with both the plant species and the transformation method employed. Challenges arise in ensuring these markers are effective and efficient, requiring careful optimization of transformation protocols and selection conditions to maintain high transformation efficiency. Only a limited number of marker genes, such as hygromycin phosphotransferase (hpt), neomycin phosphotransferase II (nptII), and bar, are commonly used in plant research and crop development, and no single selectable marker gene has been found to be universally effective in all situations [5,6]. To address this challenge, we recently used a split selectable marker system to facilitate gene stacking in plants [7]. This system contains two independent binary vectors, each carrying distinct genes of interest. Unlike the traditional co-transformation system, in which each vector typically contains a different selectable marker, each vector in our split selectable marker system contains a partial fragment of the selectable marker gene, which is connected to a partial intein fragment. The partial intein fragment refers to a segment of an intein that facilitates protein splicing in engineered systems. After the two vectors are co-transformed into the same plant cell, intein-mediated protein splicing occurs post-translationally, resulting in the reassembly of two marker protein fragments into a full-length selectable marker protein. In this natural process, an intein, a protein segment within a precursor protein, self-excises and ligates the remaining protein fragments (exteins) to form a functional protein [8]. This innovative system simplifies the identification of transgenic events generated by co-transformation using a single selection agent (e.g., antibiotic). This approach avoids the inefficiencies and incompatibilities associated with using two different selectable markers in the traditional co-transformation. Designing and constructing intein-mediated selectable markers is highly relevant to molecular cloning. Here, we describe in great detail the methodology—used in the original publication by Yuan et al. [7]—for creating split-marker constructs, co-transforming two vectors in two plant species (Arabidopsis thaliana and Populus tremula × P. alba clone INRA 717-1B4), and confirming transgenic events. Also, we created ready-to-use binary vectors to simplify the cloning procedure for creating DNA constructs for co-transformation.
Materials and reagents
Reagents
1. Agar (PhytoTech LABS, catalog number: HWW1000003C)
2. Agarose (GenBiotech, catalog number: RU1010)
3. Acetosyringone (Sigma-Aldrich, catalog number: D134406)
4. Cefotaxime (PhytoTech LABS, catalog number: C380)
5. NEBridge® Golden Gate Assembly kit (BsaI-HF® v2) (NEB, catalog number: E1601L)
6. NEB® 5-alpha competent E. coli (NEB, catalog number: C2987H)
7. PCR Purification kit (Inbio Highway, catalog number: K1206)
8. EHA105 Agrobacterium ElectroCompetent cells
9. Plasmid DNA Purification kit (Qiagen, catalog number: 12123)
10. Restriction enzymes: Depending on the restriction sites on the inserts that the user cloned
11. Sterilizing agents: ethanol
12. Silwet L-77
13. GoTaq® G2 Master Mixes (Promega, catalog number: 0000434787)
14. Murashige & Skoog basal medium with vitamins (MS) (PhytoTech LABS, catalog number: M5531)
15. MES (Sigma, catalog number: SLCF3242)
16. NAA (Sigma, catalog number: RNBJ597610)
17. IBA (PhytoTech LABS, catalog number: I538)
18. BAP (PhytoTech LABS, catalog number: B130)
19. 2iP; 6-(γ,γ-Dimethylallylamino)purine Solution (PhytoTech LABS, catalog number: D217)
20. 0.5 M EDTA (Millipore, catalog number: 3740903)
21. 5 M NaCl (Sigma, catalog number: SLBR7232V)
22. 1 M Tris-HCl (Invitrogen, catalog number:15567-027)
23. Chloroform, EMSURE® ACS, ISO, Reag. Ph. (VWR, catalog number: 1.02445.1000)
24. Isopropanol (2-Propanol) (VWR, catalog number: 1.09634.5000)
25. L-Glutamine (PhytoTech LABS, catalog number: HWY0229022A)
26. Phyto agar (Sigma, catalog number: P8169)
27. Kanamycin (PhytoTech LABS, catalog number: HHW47510054)
28. LB broth with agar (Sigma, catalog number: L3147)
29. LB broth (Sigma, catalog number: L3022)
30. Sucrose (Cicarelli, catalog number: 841214)
31. Sterile water
32. Timentin (GoloBio, catalog number: T-104-25)
33. Liquid nitrogen
34. Gel Extraction kit (Zymoclean Gel DNA Recovery kit, catalog number: D4008)
35. Q5® High-Fidelity 2× master mix (NEB, catalog number: M0492S)
36. TE buffer (Thermo Fisher Scientific, catalog number: 12090015)
37. Thidiazuron (TDZ) solution, 1 mg/mL (PhytoTech Labs, catalog number: T7999)
38. Cetyltrimethylammonium bromide (CTAB) (Sigma-Aldrich, catalog number: 219374)
Primers (for genotyping)
eYGFPuv_F: 5'-CACGGCAACCTCAACG-3'
eYGFPuv_R: 5'-CTCGACACGTCTGTGGG-3'
Solutions
1. Dip solution (see Recipes)
2. Callus induction media (CIM) (see Recipes)
3. Shoot induction media (SIM) (see Recipes)
4. Shoot elongation media (SEM) (see Recipes)
5. Root induction media (RM) (see Recipes)
6. LB agar medium (see Recipes)
7. LB medium (see Recipes)
8. MS induction medium (see Recipes)
9. 3% CTAB extraction buffer (see Recipes)
Recipes
1. Dip solution
5% sucrose
0.03% Silwet L-77
120 mL deionized water
2. Callus induction media (CIM)
4.3 g/L MS with vitamins
30 g/L sucrose
3 g/L Phyto agar
0.2 g/L L-glutamine
0.25 g/L MES
1 mL of 10 μM/L NAA
Sterilize and add 2ip (5 μM final) 1 mL of 5 mM 2ip stock, timentin (200 mg/mL stock) 1 mL/L, cefotaxime (300 mg/mL stock) 1 mL/L, and kanamycin at 100 mg/L (final concentration).
3. Shoot induction media (SIM)
4.3 g/L MS with vitamins
30 g/L sucrose
3 g/L Phyto agar
0.2 g/L L-glutamine
0.25 g/L MES
50 μL of 1 mg/mL TDZ
Sterilize and add timentin (200 mg/mL stock) 1 mL/L, cefotaxime (300 mg/mL stock) 1 mL/L, and kanamycin at 100 mg/L (final concentration).
4. Shoot elongation media (SEM)
4.3 g/L MS with vitamins
30 g/L sucrose
3 g/L Phyto agar
0.2 g/L L-glutamine
0.25 g/L MES
100 μL of 1 mg/mL BAP
Sterilize and add timentin (200 mg/mL stock) 1 mL/L, cefotaxime (300 mg/mL stock) 1 mL/L, and kanamycin at 100 mg/L (final concentration).
5. Root induction media (RM)
2.15 g/L MS with vitamins
20 g/L sucrose
3 g/L Phyto agar
0.2 g/L L-glutamine
0.25 g/L MES
100 μL of 1 mg/mL IBA
Sterilize and add timentin (200 mg/mL stock) 1 mL/L, cefotaxime (300 mg/mL stock) 1 mL/L, and kanamycin at 100 mg/L (final concentration).
6. LB agar medium
40 g/L LB broth with agar
7. LB medium
20 g/L LB broth
8. MS induction medium
4.43 g/L MS with vitamins
20 g/L sucrose
0.5 g/L MES
20 μM (final concentration) acetosyringone
9. 3% CTAB extraction buffer (50 mL)
Dissolve 1.5 g of CTAB in 16.5 mL of distilled water on a heating plate.
Add 7.5 mL of 1 M Tris-HCl (pH 7.4), 26 mL of 5 M NaCl, and 0.2 mL of 0.5 M EDTA (pH 8).
Filter-sterilize using a 0.2 μm filter.
Laboratory supplies
1. 200 μL PCR tubes (Sigma, catalog number: Z316121)
2. 1.5 mL microcentrifuge tubes (Sigma, catalog number: SLMTBP15-EP)
3. 50 mL Falcon tubes (VWR, catalog number: 21008-940)
4. 14 mL round-bottom tubes (Sigma, catalog number: Z617806)
5. Plant culture vessel (PhytoCon, 16 oz, 473 mL)
6. Pipette and pipette tips (Eppendorf)
7. 100 mm × 15 mm Petri dishes (Sigma, catalog number: Z666246)
8. Scalpel and forceps
9. Gloves
Equipment
1. E. coli and Agrobacterium pulser transformation apparatus (Bio-Rad, model: 155103)
2. High-speed centrifuges (Thermo Fisher Scientific, SorvallTM ST8 Small Benchtop Centrifuge; catalog number: 75007200)
3. Floor model centrifuge (Thermo Fisher Scientific, SorvallTM ST8 FR Floor-Standing Refrigerated Centrifuge; catalog number: 75007208)
4. Incubator (Sigma, catalog number: Z763330)
5. Incubator shaker (Sigma, model: Innova 44 incubator shaker)
6. Biosafety cabinets (LABCONCO, catalog number: 302380011)
7. Plant growth chamber (NORLAKE Scientific)
8. Autoclave
9. Vortex (FISHER MINI VORTEXER, catalog number: 02215365)
10. Nanodrop (Denovix DS11)
11. Gel Doc (Azure Biosystems 600)
12. Cuvette (Bio-Rad, 0.2 cm electrode gap cuvette, catalog number: 1652086)
Software and datasets
1. SnapGene (GSL Biotech LLC, https://www.snapgene.com/)
Procedure
A. Design and construction of vectors for plant co-transformation
1. Construction of vectors 1 and 2 using Golden Gate Assembly
a. Vectors pSplit 1 and pSplit 2 were modified as destination vectors, ready for Golden Gate Assembly.
b. Combine the destination vector pSplit 1 with insert 1 and 2 and mix the destination vector pSplit 2 with insert 3 and 4 (Figure 1) (see General note 1).
c. Assemble all fragments according to the user manual provided by NEB (see General note 2).
Figure 1. Vector cloning using Golden Gate Assembly. Inserts 1 and 2 contain Gene 1 and Gene 2, respectively, while Inserts 3 and 4 contain Gene 3 and Gene 4, respectively. pSplit 1 serves as the vector backbone and is mixed with Inserts 1 and 2 to assemble into Vector 1 via Golden Gate Assembly. Similarly, pSplit 2 serves as the vector backbone and is mixed with Inserts 3 and 4 to assemble into Vector 2 via Golden Gate Assembly. Although two inserts are used as examples for each vector, additional inserts can also be efficiently assembled using this method.
2. Gene sequence selection and primer design
a. Identify the gene sequences to be inserted into the vectors and their promoters and terminators (see General note 3).
b. Design primers for PCR amplification of the gene sequences, including restriction sites (BsaI) for cloning (Figure 2).
Figure 2. Primer design for vector cloning. A. Primer design of single insert. B. Primer design of double inserts. C. Primer design of three inserts. Red sequences indicate BsaI restriction sites. Underlined sequences represent the 4-bp overhangs required for Golden Gate Assembly. The sequences NNN---NNN indicate the annealing sequence with the target sequence.
3. PCR amplification
a. Perform PCR to amplify the target gene sequences using Q5 high-fidelity DNA polymerase (see General note 4).
b. Verify the PCR products by running a sample on an agarose gel.
c. Purify the PCR product using a gel extraction kit.
4. Transformation into E. coli
a. Transform the ligated products into competent E. coli cells using heat shock following the manufacturer’s protocol for C2987H.
b. Plate the transformed cells on LB agar plates containing kanamycin (50 μg/mL).
c. Incubate the plates overnight at 37 °C.
5. Colony PCR screening
a. Pick individual colonies for colony PCR using GoTaq® G2 Master Mixes.
b. Verify the PCR products by running a sample on an agarose gel.
c. Inoculate the positive colonies into liquid LB medium containing kanamycin (50 μg/mL).
d. Incubate the culture tubes overnight at 37 °C.
6. Plasmid Miniprep
a. Isolate plasmid DNA from the overnight cultures using a miniprep kit.
b. Verify the presence and orientation of the inserts by restriction digestion (see General note 5).
7. Plasmid sequencing
a. Sequence the purified plasmid DNA to confirm the correct insertion and orientation of the gene sequences.
b. Use sequencing primers that anneal to the plasmid backbone flanking the insert.
B. Agrobacterium transformation
1. Thaw the electrocompetent Agrobacterium EHA105 on ice.
2. Add 1 μL of constructed plasmid containing pSplit1 and pSplit2 into Agrobacterium separately and mix well by gently flicking.
3. Transfer the mixture immediately to the electroporation cuvette.
4. Electroporate at 2.4 kV until the apparatus beeps.
5. Add 1 mL of LB medium into the cuvette, resuspend, and transfer back to an Eppendorf tube.
6. Incubate for 1–2 h at 30 °C with shaking at 180 rpm.
7. Plate on LB agar plates with kanamycin (50 μg/mL) and rifampicin (50 μg/mL).
8. Incubate the plates for 48 h at 30 °C.
9. Pick individual colonies for colony PCR using GoTaq® G2 Master Mixes.
10. Verify the PCR products by running a sample on an agarose gel.
C. Agrobacterium-mediated co-transformation in Arabidopsis thaliana
1. Preparation of Agrobacterium tumefaciens cultures
a. Start a preculture by inoculating each of two selected Agrobacterium tumefaciens strains containing pSplit1 or pSplit2 into 5 mL of LB medium with kanamycin (50 μg/mL) and rifampicin (50 μg/mL) in a 50 mL Falcon tube. (see General note 6).
b. Incubate the culture overnight at 28 °C with shaking at 180 rpm and monitor growth. Alternatively, monitor for two nights if necessary until OD reaches 0.8–1.0 (see General note 7).
c. Inoculate 10 μL of the preculture into 100 mL of LB medium with antibiotic selection in a sterile Erlenmeyer flask.
d. Incubate overnight at 28 °C with shaking at 180 rpm.
2. Dipping of the plants
a. Verify that both cultures have grown well (see General note 7).
b. Prepare the floral dip solution (see Recipes).
c. For each cultured strain, divide the 100 mL culture into two Falcon tubes.
d. Centrifuge at 3,500× g for 7–10 min.
e. Carefully remove the supernatant.
f. Resuspend the pellet in 30 mL (total volume) of LB medium without antibiotics.
g. Pour 120 mL of floral dip solution into a plastic-covered tray and add 30 mL of bacterial suspension from each of the two strains (a total of 60 mL) to the center of the dip solution.
h. Submerge each plant completely in the liquid for a few seconds (see General note 8).
i. Place the dipped plants back in the original pots or transfer them to an appropriate container.
j. Dip all the plants of the same construct and then clean up the area.
k. Leave the plants on the bench for 48 h.
l. Transfer the plants to the growth chamber (temperature: 22–25 °C; photoperiod: 16/8 h light/dark cycle; humidity: 50%–55%).
m. Once plants are dry, you can collect the seeds.
D. Agrobacterium-mediated co-transformation in poplar (Populus tremula × P. alba clone INRA 717-1B4)
1. Preparation of Agrobacterium tumefaciens cultures
a. Streak Agrobacterium tumefaciens (EHA105) of two constructs on an LB agar plate containing kanamycin (50 μg/mL) and rifampicin (50 μg/mL). Then, incubate the plate at 28 °C for 2 days.
b. Inoculate a single A. tumefaciens colony into 5 mL of LB medium with kanamycin (50 μg/mL) and rifampicin (50 μg/mL) and grow overnight at 28 °C with shaking (180 rpm).
c. Add 100 µL of culture into 50 mL of LB medium with kanamycin (50 μg/mL) and rifampicin (50 μg/mL) and grow overnight at 28 °C with shaking (180 rpm).
d. The next day, centrifuge the cultures at 3,500× g for 15 min at room temperature to pellet the cells.
e. Discard the supernatant and proceed to the next step.
2. Resuspension of Agrobacterium pellets
a. Resuspend each Agrobacterium pellet in MS induction medium.
b. Ensure the MS induction medium contains 20 μM acetosyringone to enhance virulence.
c. Adjust the OD600nm of the suspension to 0.5 using the MS induction medium.
3. Preparation of poplar leaf disks
a. Excise approximately 150 young leaf disks (~0.5 cm diameter) from healthy poplar 717 (Populus tremula × alba clone INRA 717-1B4) leaves using a sterilized punch or scalpel (see General note 9).
b. Keep the leaf disks on sterile filter paper moistened with MS medium until ready for infection.
4. Agrobacterium infection
a. Mix an equal volume of MS induction medium containing two split constructs into a 50 mL conical flask.
b. Soak the excised leaf disks in the Agrobacterium suspension for 1 h.
c. Agitate gently during the infection process to ensure even exposure.
5. Co-culture
a. Transfer the soaked leaf disks onto solid co-culture medium (e.g., MS medium supplemented with the required hormones and 20 μM acetosyringone).
b. Incubate the plates in the dark at 22–25 °C for 2–3 days to allow co-culturing.
6. Washing
a. After co-culture, wash the leaf disks thoroughly with sterile water containing antibiotics (e.g., 300 mg/L cefotaxime and 200 mg/L timentin) for 1 h to remove excess Agrobacterium.
b. Blot the disks dry on sterile filter paper before transferring them to the callus induction medium (CIM) containing antibiotics (e.g., 300 mg/L cefotaxime, 200 mg/L timentin, and 100 mg/L kanamycin).
7. Callus induction
a. Transfer the washed leaf disks onto solid callus induction medium (e.g., MS medium supplemented with appropriate plant growth regulators (see CIM recipe).
b. Incubate the plates under controlled light conditions (16/8 h light/dark cycle) at 22–25 °C.
c. Monitor the disks for callus formation, which may take 2–4 weeks.
d. Subculture immediately if Agrobacterium re-growth is observed.
8. Shoot induction
a. Transfer the developing calli to shoot induction medium (SIM) (e.g., MS medium with cytokinin and antibiotics; see Recipes).
b. Continue incubation under the same light and temperature conditions until shoots begin to emerge.
c. Subculture every 2–3 weeks. Adventitious shoots typically emerge within 4–6 weeks. Separate transformation events as early as possible during subculturing, either at the callus or multiple shoot stages.
9. Shoot elongation
a. Transfer emerging shoots to shoot elongation medium (SEM) (e.g., MS medium with a lower concentration of cytokinin and antibiotics; see Recipes).
b. Incubate until the shoots have elongated sufficiently, which typically takes 2–3 weeks.
10. Root induction
a. Transfer elongated shoots to root induction medium (RM) (e.g., MS medium with auxin).
b. Incubate under appropriate conditions until roots develop, confirming successful transformation.
E. Confirmation of transformation
1. Sample collection: collect leaf samples (0.5–1.0 cm) from the Arabidopsis and poplar 717 lines (see General note 10).
2. Leaf grinding
a. Place the collected leaf samples in a pre-cooled mortar.
b. Grind the leaves into a fine powder using liquid nitrogen.
3. Genomic DNA isolation
a. Add 500 μL of 3% CTAB extraction buffer to 100 mg of powdered plant material in a 2.0 mL microcentrifuge tube. Vortex the mixture vigorously to ensure thorough mixing.
b. Incubate the tubes in a 65 °C water bath for 30 min. During the incubation, invert the tubes every 5–10 min to keep the contents mixed.
c. After incubation, centrifuge the samples at 13,000× g for 10 min. Carefully transfer the supernatant to a new 1.5 mL microcentrifuge tube.
d. Add 600 μL of chloroform to the supernatant and vortex thoroughly.
e. Centrifuge at 13,000× g for 10 min. Transfer the upper aqueous phase to a clean 1.5 mL microcentrifuge tube.
f. Add an equal volume of chloroform to the transferred aqueous phase and vortex to mix.
g. Centrifuge the mixture at 13,000× g for 10 min. Again, transfer the upper aqueous phase to a clean 1.5 mL tube.
h. Add an equal volume of isopropanol (2-propanol) to the aqueous phase. Mix by pipetting up and down, then precipitate the DNA by placing the tube at -20 °C for 30 min.
i. Centrifuge the precipitated mixture at 13,000× g for 20 min. Carefully discard the supernatant, avoiding disturbance of the DNA pellet, and allow the pellet to air dry briefly.
j. Add approximately 500 μL of chilled 70% ethanol to the pellet and centrifuge at 13,000× g for 5 min to wash the DNA pellet. Discard the ethanol and allow the pellet to air dry completely.
k. Finally, dissolve the DNA pellet in 100 μL of 1× TE buffer.
4. PCR setup
PCR reaction mixture (25 μL volume)
GoTaq® G2 Green master mix, 2× 12.5 μL
eYGFPuv_F (10 μM) 2.5 μL
eYGFPuv_R (10 μM) 2.5 μL
DNA template 1.0 μL
Nuclease-free water to 25 μL
a. For eYGFPuv genotyping, add the eYGFPuv_F (5'-CACGGCAACCTCAACG-3') and eYGFPuv_R (5'-CTCGACACGTCTGTGGG-3') (see General note 11).
b. Mix the reagents gently.
5. PCR cycling conditions (Table 1):
Table 1. PCR cycling conditions
Step Temperature Time Cycle
Initial denaturation 95 °C 2 min 1 cycle
Denaturation 95 °C 10–30 s 25–35 cycles
Annealing 55–60 °C 30 s
Extension 72 °C 1 min/kb
Final extension 72 °C 5 min 1 cycle
Rest 4 °C Indefinite 1 cycle
6. Post-PCR analysis
Analyze the PCR products using agarose gel electrophoresis to confirm the presence of the target genes.
Validation of protocol
This protocol was used to generate the co-overexpression lines by Yuan et al. [7].
General notes and troubleshooting
General notes
1. The number of DNA fragments varies based on user needs, with successful assemblies reported ranging from 1 to 16 fragments.
2. Golden Gate Assembly protocol for using NEBridge® Golden Gate Assembly Kit (BsaI-HF®v2) (E1601) can be found at https://www.neb.com/en-us/protocols/2018/10/02/golden-gate-assembly-protocol-for-using-neb-golden-gate-assembly-mix-e1601. The manual does not mention that the destination plasmid requires pre-digestion with the BsaI enzyme followed by gel purification. However, we found that performing pre-digestion and gel purification significantly improves assembly efficiency, especially when the number of inserts exceeds two.
3. For gene overexpression, the gene sequence can either be the coding sequence of the gene of interest (without introns) or its genomic sequence (containing introns), as both are effective.
4. DNA fragments to be inserted into the binary vectors can also be commercially synthesized by vendors such as Integrated DNA Technologies (IDT, Coralville, IA) and Twist Bioscience (South San Francisco, CA). The amount of a DNA insert varies depending on the user’s purpose. Single or multiple gene cassettes can be inserted into destination plasmids through Golden Gate Assembly in a one-pot reaction.
5. Restriction digestion is optional here if positive colonies have been previously verified through colony PCR.
6. Only use a fresh colony. Do not close the Falcon tube completely and tape the lid with micropore tape.
7. The culture must be well-grown and clearly turbid.
8. Ensure the rosette is submerged during floral dipping for Arabidopsis transformation. You will observe air bubbles between the leaves and stem.
9. To achieve optimal transformation efficiency, avoid using overly mature leaves. Instead, select leaves from 1–2-month-old plants grown in vitro.
10. We detected two co-transformed vectors in both the T1 and T2 generations of Arabidopsis. Kanamycin (50 μg/mL) was used during seed germination to select for successful transformation.
11. You may design specific primers tailored to your gene of interest.
Acknowledgments
Funding: The work was supported by the Center for Bioenergy Innovation (CBI), a DOE Research Center supported by the Biological and Environmental Research (BER) program. Oak Ridge National Laboratory is managed by UT-Battelle, LLC for the U.S. DOE under Contract Number DE-AC05-00OR22725. The United States Government retains and the publisher, by accepting the article for publication, acknowledges that the United States Government retains a non-exclusive, paid-up, irrevocable, worldwide license to publish or reproduce the published form of this manuscript or allow others to do so, for United States Government purposes. This protocol has been validated in Yuan et al. [7].
Competing interests
The authors declare no conflict of interest.
References
Roberts, D. and Mattoo, A. (2018). Sustainable Agriculture—Enhancing Environmental Benefits, Food Nutritional Quality and Building Crop Resilience to Abiotic and Biotic Stresses. Agriculture. 8(1): 8.
Naqvi, S., Farré, G., Sanahuja, G., Capell, T., Zhu, C. and Christou, P. (2010). When more is better: multigene engineering in plants. Trends Plant Sci. 15(1): 48–56.
Ceccon, C. C., Caverzan, A., Margis, R., Salvadori, J. R. and Grando, M. F. (2020). Gene stacking as a strategy to confer characteristics of agronomic importance in plants by genetic engineering. Ciência Rural. 50(6): e20190207.
Halpin, C. (2005). Gene stacking in transgenic plants – the challenge for 21st century plant biotechnology. Plant Biotechnol J. 3(2): 141–155.
Miki, B. and McHugh, S. (2004). Selectable marker genes in transgenic plants: applications, alternatives and biosafety. J Biotechnol. 107(3): 193–232.
Ahmed, S., Hulbert, A. K., Xin, X. and Neff, M. M. (2024). The ability of Arabidopsis to recover from Basta and its application in isolating Cas9-free mutants. Front Plant Sci. 15: e1408230.
Yuan, G., Lu, H., De, K., Hassan, M. M., Liu, Y., Islam, M. T., Muchero, W., Tuskan, G. A. and Yang, X. (2023). Split selectable marker systems utilizing inteins facilitate gene stacking in plants. Commun Biol. 6(1): 567.
Perler, F. B. (2002). InBase: the Intein Database. Nucleic Acids Res. 30(1): 383–384.
Article Information
Publication history
Received: Sep 3, 2024
Accepted: Jan 5, 2025
Available online: Jan 16, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Category
Plant Science > Plant transformation > Agrobacterium
Biological Sciences > Biological techniques
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5,216 | https://bio-protocol.org/en/bpdetail?id=5216&type=0 | # Bio-Protocol Content
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Peer-reviewed
Generation, Propagation, and Titering of Dicistrovirus From an Infectious Clone
JS Junzhou Shen
Jibin Sadasivan
EJ Eric Jan
In Press, Available online: Jan 22, 2025
DOI: 10.21769/BioProtoc.5216 Views: 26
Reviewed by: Keisuke TabataChhuttan L Meena Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in PLOS Pathogens Dec 2022
Abstract
Cricket paralysis virus (CrPV), a member of the family Dicistroviridae, is a single-stranded positive-sense RNA virus that primarily infects arthropods. Some members of the dicistrovirus family, including the honey bee viruses Israeli acute paralysis virus and Acute bee paralysis virus and the shrimp-infecting Taura syndrome virus, pose significant threats to agricultural ecosystems and economies worldwide. Dicistrovirus infection in Drosophila is used as a model system to study fundamental insect–virus–host interactions. The availability of a CrPV infectious clone allows controlled manipulation of the viral genome at a molecular level. Effective viral propagation and titration techniques are crucial for understanding the pathogenesis and epidemiology of dicistrovirus infections. Traditional methods for assessing viral titers, such as plaque assays, are unsuitable for CrPV, since Drosophila tissue culture cells like Schneider 2 cells cannot readily form adherent plaques. Here, we present a streamlined protocol for generating a recombinant virus from a CrPV infectious clone, propagating the virus in S2 cells and titering the virus by an immunofluorescence-based focus-forming assay (FFA). This protocol offers a rapid and reliable approach for generating recombinant viruses, viral amplification, and determining CrPV titers, enabling efficient investigation into viral biology and facilitating the development of antiviral strategies.
Key features
• Generate recombinant virus from infectious clones.
• Sequential amplification protocol for scalable virus production.
• Repeated freeze-thawing for virus harvesting.
• Immunostaining focus-forming assay (FFA) for CrPV titration.
• Focus-forming units (FFU) quantified using a high-throughput microscopic screening platform.
Keywords: Cricket paralysis virus Infectious clone Amplification Dicistrovirus Infection
Graphical overview
Graphical overview of cricket paralysis virus (CrPV) infectious clone generation, propagation, sequence confirmation, titering, and infection. A. CrPV infectious clone preparation and propagation. CrPV cDNA plasmid is amplified via bacterial transformation, linearized by restriction enzyme digestion, and used in an in vitro transcription reaction to produce CrPV genomic RNA. S2 cells are transfected with the CrPV RNA and then incubated for 48 h to allow viral protein synthesis and viral replication. Infected cells are harvested and subjected to three freeze-thaw cycles to release intracellular virus, and the viral stock is aliquoted and stored at -80 °C. B. Propagation of CrPV in S2 cells via sequential amplification. S2 cells are infected with CrPV and incubated for 18–24 h. After incubation, cells are harvested and subjected to three freeze-thaw cycles to release the virus. The harvested virus is used to reinfect fresh S2 cells in a sequential amplification step to increase viral yield. The virus is collected from the supernatant and aliquoted for storage at -80 °C for further experiments. C. CrPV genome validation. Viral RNA is purified from aliquots of CrPV, reverse transcribed, and then PCR amplified using CrPV-specific primers. The PCR product is analyzed by DNA gel electrophoresis and verified by Sanger sequencing. D. Titering CrPV. Serial dilutions of the virus (10-fold dilution) are used to infect S2 cells for 6 h and then immunostained with anti-CrPV capsid protein (VP2) antibody followed by secondary staining using Texas red-conjugated antibodies. The infected cells are visualized and quantified using a high-throughput microscopic screening platform. The viral titer is calculated based on the number of VP2-positive cells. D. RNA gel electrophoresis and western blotting for infection validation. S2 cells are infected with CrPV at specific multiplicity of infection (MOI) values and harvested after defined time points. RNA is extracted from the infected cells and analyzed via RNA gel electrophoresis and western blotting targeting CrPV-1A and CrPV-VP2 to assess the progression of the infection.
Background
Cricket paralysis virus (CrPV), a small RNA virus from the Dicistroviridae family, primarily infects arthropods, including economically significant pests and the model organism Drosophila melanogaster [1,2]. The genome organization of CrPV includes two internal ribosome entry sites (IRESs) that direct the translation of non-structural and structural protein-encoding open reading frames (ORF1 and ORF2), as shown in the schematic representation of the CrPV genome (Figure 1) [2]. The availability of an infectious CrPV clone has proven invaluable for introducing precise mutations into viral RNA and proteins, allowing researchers to investigate the specific roles of individual viral components during infection [3–6]. As CrPV serves as a model system for studying fundamental virus–host interactions, the development of efficient methods for propagating and quantifying the virus is critical. Traditional viral titration methods, such as plaque assays, involve counting plaques formed by the lysis of virus-infected cells. However, Drosophila S2 cells are semi-adherent and cannot form the dense monolayers required for plaque formation by CrPV. As a result, CrPV-infected cells fail to produce clear, countable plaques, making them unsuitable for standard plaque assays. This limitation highlights the need for alternative methods to accurately and efficiently quantify CrPV.
Our protocol for CrPV propagation and titration addresses these challenges. A sequential amplification approach is used, wherein an initial infection of Drosophila S2 cells (30 million) is performed. Subsequently, the intracellular virus is harvested via freeze-thaw cycles and used to infect a larger batch of cells (100 million) to amplify the virus. This step helps produce higher viral yields by harvesting the cells before lysis. Finally, infected cells are then subject to freeze-thaw to disrupt cellular membranes to release the accumulated intracellular viral particles. This method helps with the continuous amplification of the virus.
For viral titration, we use an immunostaining-based focus-forming assay (FFA), a limiting dilution assay that quantifies the number of infectious particles in a given volume. Viral stocks are serially diluted and used to infect S2 cells in a 96-well plate that has been pre-treated with Concanavalin A to enhance cell adherence [7]. After incubation, cells are fixed with paraformaldehyde, permeabilized with methanol, and stained with antibodies against the CrPV capsid protein, VP2. The fluorescence signal shows the presence of VP2 expression within cells, serving as an indicator of the infection status. Fluorescence microscopy, such as using a high-throughput imaging system (e.g., CellInsightTM CX5 High Content Screening), is used to visualize and quantify the infected cell foci, expressed as focus-forming units (FFU) per milliliter. This immunostaining-based titration method can be effectively used to determine viral titer when plaque assay is not an option. By providing a reliable method for virus propagation and titration, our protocol advances the study of insect viruses, contributing to the broader understanding of viral biology and the development of novel antiviral strategies.
Figure 1. RNA genome of cricket paralysis virus (CrPV). CrPV is a positive-sense, single-stranded RNA. It contains two internal ribosome entry sites (IRESs) that facilitate the translation of two distinct open reading frames (ORF1 and ORF2). The 5' UTR IRES initiates translation of ORF1, which encodes non-structural proteins involved in viral replication, including 1A, 2B, 2C, 3A, 3B, 3C, and the RNA-dependent RNA polymerase (RdRp or 3D), as well as four viral protein genomes (VPgs). The intergenic region (IGR) IRES initiates the translation of ORF2, which encodes structural proteins essential for forming the viral capsid: VP2, VP3, VP1, and VP4. The CrPV genome is flanked by a 5' UTR with an IRES structure and a 3' UTR that ends with a poly(A) tail, which are both crucial for viral replication and translation efficiency. This figure is adapted from Warsaba et al. [2], Dicistrovirus-Host Molecular Interactions, Current Issues in Molecular Biology.
Materials and reagents
Biological materials
1. Cricket paralysis virus (CrPV) infectious clone, generated based on the methodology described in the Journal of Virology article titled "The 5' Untranslated Region of a Novel Infectious Molecular Clone of the Dicistrovirus Cricket Paralysis Virus Modulates Infection" by Kerr et al. [3]
2. 5-alpha competent E. coli (New England Biolabs, catalog number: C2987)
3. Drosophila Schneider 2 (S2) cells cultured in Schneider's Drosophila medium (Thermo Fisher Scientific, catalog numbers: R69007)
Reagents
1. 1 kb plus DNA ladder (New England Biolabs, catalog number: N3232)
2. 100 bp DNA ladder (New England Biolabs, catalog number: N3231)
3. Acetic acid (Sigma-Aldrich, catalog number: 45754)
4. Agar (Sigma-Aldrich, catalog number: 05040)
5. Ammonium persulfate (APS) (Sigma-Aldrich, catalog number: A3678)
6. Ampicillin sodium salt (Sigma-Aldrich, catalog number: A8351)
7. α-CrPV-1A antibody (custom protein antibody, Genscript) [5]
8. α-CrPV-VP2 antibody (custom peptide antibody, Genscript – C-ATFQDKQENSHIENE-NH2 and C-KLWIHKTYLKRPAR-NH2) [8]
9. Bleach/sodium hypochlorite solution (Sigma-Aldrich, catalog number: 1.05614)
10. Blue protein loading dye (New England Biolabs, catalog number: B7703)
11. Bovine serum albumin (BSA) (Sigma-Aldrich, catalog number: A7906)
12. Bradford protein assay dye reagent concentrate (Bio-Rad, catalog number: 5000006)
13. Chemiluminescence kit (Thermo Fisher Scientific, catalog number: 89880)
14. Chloroform (Thermo Fisher Scientific, catalog number: 043685.M6)
15. Concanavalin A (Con A) (Sigma-Aldrich, catalog number: 11028-71-0)
16. Diethyl pyrocarbonate (DEPC) (Sigma-Aldrich, catalog number: D5758)
17. Donkey anti-rabbit IgG-horseradish peroxidase (Amersham, catalog number: NA934)
18. DNase I (New England Biolabs, catalog number: M0303)
19. Ecl136II restriction enzyme (Thermo Fisher Scientific, catalog number: ER0251)
20. EDTA (ethylenediaminetetraacetic acid, disodium salt) (Sigma-Aldrich, catalog number: 798681)
21. Ethanol (Sigma-Aldrich, catalog number: 1.00974)
22. Fetal bovine serum (FBS) (Gibco, catalog number: 16-000-069)
23. Gel loading dye purple (6×) (New England Biolabs, catalog number: B7024)
24. Glacial acetic acid (Sigma-Aldrich, catalog number: 1005706)
25. Glycerol (Sigma-Aldrich, catalog number: G7893)
26. Goat anti-mouse IgG-horseradish peroxidase (Santa Cruz Biotechnology, catalog number: sc-2005)
27. Guanidine hydrochloride (Sigma-Aldrich, catalog number: G3272)
28. Halt protease and phosphatase inhibitor cocktail (100×) (Thermo Fisher Scientific, catalog number: 78444)
29. Hoechst (Invitrogen, catalog number: H21486)
30. IGEPAL CA-630 (Nonidet P-40, NP-40 substitute) (Sigma-Aldrich, catalog number: 56741)
31. Isopropanol (Thermo Fisher Scientific, catalog number: 040983.M1)
32. Lipofectamine 2000 transfection reagent (Thermo Fisher Scientific, catalog number: 11668027)
33. LunaScript RT supermix kit (New England Biolabs, catalog number: E3010)
34. Methanol (Fisher Scientific, catalog number: 67-56-1)
35. Milk powder (Sigma-Aldrich, catalog number: 1443825)
36. Molecular-grade agarose (FroggaBio, catalog number: A87-500G)
37. Monarch PCR & DNA Cleanup kit (5 μg) (New England Biolabs, catalog number: T1030)
38. MOPS [3-(N-morpholino)propanesulfonic acid] (Sigma-Aldrich, catalog number: 475898)
39. Gibco Opti-MEM reduced serum medium (Fisher Scientific, catalog number: 31985070)
40. Paraformaldehyde (Fisher Scientific, catalog number: AC416785000)
41. Penicillin G potassium salt (Sigma-Aldrich, catalog number: 113-98-4)
42. Phusion High-Fidelity PCR kit (New England Biolabs, catalog number: E0553)
43. Potassium chloride (Sigma-Aldrich, catalog number: 529552)
44. Potassium phosphate monobasic (Sigma-Aldrich, catalog number: 529568)
45. Qubit RNA High-Sensitivity (HS) Assay kit (Invitrogen, catalog number: Q32855)
46. Riboruler RNA ladder (Thermo Fisher Scientific, catalog number: SM1821)
47. RiboLock RNase inhibitor (Thermo Fisher Scientific, catalog number: EO0382)
48. Ribonucleotide solution mix (New England Biolabs, catalog number: N0466)
49. RNA Clean-Up kit (New England Biolabs, catalog number: T2040)
50. SafeView DNA stains (Applied Biological Materials, catalog number: G108)
51. SalI restriction enzyme (New England Biolabs, catalog number: R0138)
52. Schneider's Drosophila medium (Gibco, catalog number: 21720024)
53. Sodium chloride (Sigma-Aldrich, catalog number: S9888)
54. Sodium deoxycholate (Sigma-Aldrich, catalog number: 264103)
55. Sodium dodecyl sulfate (SDS) (Sigma-Aldrich, catalog number: 11667289001)
56. Sodium phosphate dibasic (Sigma-Aldrich, catalog number: 567547)
57. Streptomycin sulfate salt (Sigma-Aldrich, catalog number: 3810-74-0)
58. T7 RNA polymerase (New England Biolabs, catalog number: E2040S)
59. TEMED (Sigma-Aldrich, catalog number: T7024)
60. Texas red goat anti-rabbit IgG (H+L) secondary antibody (Invitrogen, catalog number: T-2767)
61. Tris base (Sigma-Aldrich, catalog number: 252859)
62. Tris hydrochloride (Sigma-Aldrich, catalog number: 10812846001)
63. Triton X-100 (Sigma-Aldrich, catalog number: X100RS)
64. Trizol (Thermo Fisher Scientific, catalog number: 15596026)
65. Tryptone (Sigma-Aldrich, catalog number: T2559)
66. Tween 20 (Sigma-Aldrich, catalog number: P1379)
67. Yeast extract (Sigma-Aldrich, catalog number: Y1625)
Solutions
1. Ampicillin stock solution (see Recipes)
2. Complete Schneider's Drosophila medium (see Recipes)
3. Con A solution (see Recipes)
4. DEPC-treated water (see Recipes)
5. LB-agar plate (see Recipes)
6. Liquid LB (see Recipes)
7. Penicillin and streptomycin solution (see Recipes)
8. LB-gar solution (see Recipes)
9. LB solution (see Recipes)
10. PB buffer (DNA purification) (see Recipes)
11. PE buffer (wash buffer) (see Recipes)
12. Phosphate buffered saline (PBS) solution (see Recipes)
13. 1× Tris-Acetate-EDTA (TAE) buffer(see Recipes)
14. Radio-immunoprecipitation assay (RIPA) buffer (see Recipes)
15. 10× MOPS (3-(N-morpholino)propanesulfonic acid) buffer (see Recipes)
16. 0.1% Tris-buffered saline with Tween 20 (TBST) buffer (see Recipes)
17. Resolving and stacking solutions for western blotting: made right before gel casting
18. Texas red secondary antibody in 5% BSA/PBS: it is light-sensitive; wrap with aluminum foil to prevent photobleaching
Recipes
1. Ampicillin stock solution
Dissolve ampicillin salts in DEPC-treated water to 100 mg/mL. This solution can be aliquoted to 1 mL per microfuge tube and stored at -20 °C.
2. Complete Schneider's Drosophila medium
Schneider's Drosophila medium includes 10% FBS and 1% penicillin and streptomycin solution. After mixing, filter the media using a bottle-top vacuum filter and store in an autoclaved glass bottle at 4 °C. It can be stored indefinitely if the bottle is unopened.
3. Con A solution
Con A in PBS (0.5 mg/mL), 3% paraformaldehyde, and 5% BSA in PBS. Filter by a syringe filter and store on ice.
4. DEPC-treated water
Treat Milli-Q water with 0.2% (v/v) DEPC and autoclave for 30 min in a glass bottle. Cool down the bottled water to room temperature before use. It can be stored indefinitely if the bottle is unopened.
5. LB-agar plate
Place a magnetic stir bar into the LB-agar solution and autoclave it in a glass bottle for 30 min at 121 °C. After autoclaving, stir the solution on a magnetic stir plate while allowing it to cool to approximately 50 °C. Once cooled, add ampicillin to a final concentration of 100 μg/mL and mix gently to ensure even distribution. Pour the LB-agar into Petri dishes, adding just enough to cover the bottom of each dish. Close the lids and allow the plates to cool and solidify overnight at room temperature.
6. Liquid LB
Autoclave LB solution for 30 min in a glass bottle and store at room temperature. Cool down the bottled water to room temperature before use.
7. Penicillin and streptomycin solution
Dissolve penicillin and streptomycin salts in DEPC-treated water to 6 mg/mL and 10 mg/mL, respectively. This solution can be aliquoted to 10 mL per 15 mL Falcon tube and stored at -20 °C.
9. LB-agar solution
Reagent Final concentration Quantity or Volume
Tryptone 1% (w/v) 10 g
Yeast extract 0.5% (w/v) 5 g
Sodium chloride 1% (w/v) 10 g
Agar 2% (w/v) 20 g
Deionized water (dH2O) n/a To 1 L
Total n/a 1 L
10. LB solution
Reagent Final concentration Quantity or Volume
Tryptone 1% (w/v) 10 g
Yeast extract 0.5% (w/v) 5 g
Sodium chloride 1% (w/v) 10 g
Deionized water (dH2O) n/a To 1 L
Total n/a 1 L
11. PB buffer (DNA purification)
Reagent Final concentration Quantity or Volume
Guanidine hydrochloride (GuHCl) 5 M 476.6 g
Isopropanol 30% (v/v) 300 mL
Tris-HCl (pH 6.6) 20 mM 2.42 g
Deionized water (dH2O) n/a To 1 L
Total n/a 1 L
12. PE buffer (wash buffer)
Reagent Final concentration Quantity or Volume
Tris-HCl 10 mM 1.21 g
Ethanol 80% (v/v) 800 mL
Deionized water (dH2O) n/a To 1 L
Total n/a 1 L
13. Phosphate buffered saline (PBS) solution
Reagent Final concentration Quantity or Volume
Sodium chloride 137 mM 8 g
Potassium chloride 2.7 mM 0.2 g
Sodium phosphate dibasic 10 mM 1.44 g
Potassium phosphate monobasic
Deionized water (dH2O)
1.8 mM
n/a
0.24 g
To 1 L
Total n/a 1 L
Note: Autoclave for 30 min in a 1 L glass bottle and store at room temperature.
14. 1× Tris-Acetate-EDTA (TAE) buffer
Reagent Final concentration Quantity or Volume
Tris base 40 mM 4.84 g
Glacial acetic acid 20 mM 1.14 mL
EDTA 1 Mm 0.372 g
Deionized water (dH2O) n/a To 1 L
Total n/a 1 L
Note: Store at room temperature.
15. RIPA buffer
Reagent Final concentration Quantity or Volume
Sodium chloride 150 mM 0.439 g
IGEPAL CA-630 1% 0.5 mL
Sodium deoxycholate 0.5% 0.25 g
SDS 0.1% 0.05 g
Glycerol 10% 5 mL
Tris-HCl (pH 8.0) Tris(hydroxymethyl)aminomethane 50 mM 0.3 g (adjust pH to 8)
Deionized water (dH2O) n/a To 50 mL
Total n/a 50 mL
Note: Store at 4 °C and preferably protected from light.
16. 10× MOPS buffer
Reagent Final concentration Quantity or Volume
MOPS 200 mM 41.86 g
Sodium acetate 50 mM 4.1 g
EDTA 10 mM 3.72 g
Deionized water (dH2O) n/a To 1 L
Total n/a 1 L
17. 0.1% Tris-buffered saline with Tween 20 (TBST) buffer
Reagent Final concentration Quantity or Volume
Tris base 20 mM 2.42 g
Sodium chloride 150 mM 8 g
Tween 20 0.1% 1 mL
Deionized water (dH2O) n/a To 1 L
Total n/a 1 L
Laboratory supplies
1. 1.5, 15, and 50 mL tube rack
2. 1.5 and 2 mL Eppendorf tubes (Thermo Fisher Scientific, catalog numbers: AM12450 and AM12475)
3. 15 and 50 mL Falcon tubes (FroggaBio, catalog numbers: TB15-500 and TB50-500)
4. 14 mL round-bottom test tubes (Sigma-Aldrich, catalog number: CLS352057)
5. 6-well TC-treated plate (Corning, catalog number: CLS353224)
6. Bottle-top vacuum filter (Corning, catalog number: CLS431097)
7. Centrifuge (Thermo Fisher Scientific, model: accuSpin 1-1R)
8. Disposable polystyrene cuvettes (Bio-Rad, catalog number: 2239955)
9. DNase/RNase-free tips (1,000, 200, 10 μL)
10. Kimwipes (Fisher Scientific, catalog number: 06-666)
11. Micropipettes (200–1,000, 20–200, 2–20, and 0.2–2 μL)
12. Multichannel micropipettes (20–200 μL)
13. PCR strip tubes (FroggaBio, catalog number: STF-A120)
14. Petri dishes (Thermo Fisher Scientific, catalog number: FB0875713)
15. Spin Miniprep columns (Qiagen, catalog number: 27115)
16. Sterile 96-well plate, surface treated (VWR, catalog number: 10062-900)
17. Sterile serological pipettes 10 mL (Fisher Scientific, catalog number: 13-678-12)
18. Sterile serological pipettes 25 mL (Fisher Scientific, catalog number: 13-678-14)
19. Syringe filter (Sigma-Aldrich, catalog number: SLMP025SS)
20. T25 and T75 TC flask (Thermo Fisher Scientific, catalog numbers: 156367 and 156499)
Equipment
1. Isotemp incubator (Thermo Fisher Scientific, catalog number: 151030513)
2. Bio-Rad Gel Doc 2000 (Bio-Rad, catalog number: 04678)
3. BioPhotometer spectrophotometer (Eppendorf, catalog number: 6131-FCP22)
4. CellInsight CX5 HCS platform (Thermo Fisher Scientific, catalog number: CX51110)
5. EVOS FLoid imaging system (Thermo Fisher Scientific, catalog number: 4471136)
6. NanoDrop spectrophotometer ND-1000 (Thermo Fisher Scientific, catalog number: 2353-30-0010)
7. Typhoon biomolecular imager (Cytiva, catalog number: 29187191)
Software and datasets
1. BioRender version 4.0 (BioRender.com, 10/2024)
2. HCS Navigator version 6.6.0 (Thermo Fisher Scientific, 12/2015)
3. ImageJ version 1.54g (National Institutes of Health, Bethesda, MD, USA, 2024)
4. Microsoft Excel for Microsoft 365 (Microsoft Corporation, 2024)
5. SnapGene version 6.0 (GSL Biotech, 2024)
Procedure
A. Standard culture and passaging of S2 cells
1. Warm complete S2 cell media to room temperature in a biological safety cabinet (BSC).
2. Ensure all solutions and equipment that contact the cells are sterile. Work inside a BSC using proper sterile techniques.
3. Check the culture flask under a microscope to confirm the cells are healthy, actively growing, and at an appropriate density for passaging (log to mid-log phase, approximately 0.5 × 10 to 5 × 10 cells/mL).
4. Tap the flask gently to dislodge the loosely adherent cells into the media.
5. Using a serological pipette, gently pipette the media up and down several times to rinse the flask surface and break up any visible cell clumps.
6. Transfer an appropriate volume of the cell suspension into a new cell culture flask to achieve the desired dilution ratio (e.g., 1:2 to 1:5 dilution).
7. Add fresh, prewarmed complete media to top up to the final volume:
a. T175 flask: Add up to 30 mL of media.
b. T75 flask: Add up to 15 mL of media.
c. T25 flask: Add up to 5 mL of media.
8. Place the flask in a 25 °C incubator without CO2. Ensure the cap is loosened slightly for gas exchange.
9. Monitor the culture regularly for signs of healthy growth and proper density.
10. Backup culture (Optional): Retain the old flask as a backup until the newly passaged cells are confirmed to be growing well and appear healthy under the microscope.
B. Generation of CrPV from an infectious clone
B1. Amplification of the CrPV cDNA plasmid via bacterial transformation
1. In a microfuge tube, thaw E. coli DH5α competent cells (50 μL per transformation reaction) on ice for 5 min.
2. Add 1 μL of plasmid DNA (approximately 100 ng) to the competent cells. The CrPV cDNA plasmid (pCrPV-2 or -3) is described in Kerr et al. [3].
3. Mix by gently pipetting the DNA through the cells while moving the pipette tip through the suspension. Tap tubes gently to mix.
4. Incubate the cells on ice for 30 min to allow DNA uptake.
5. Heat shock the cells at 42 °C for 45 s. Do not shake during this step.
6. Immediately transfer the cells on ice after heat shock.
7. Add 200 μL of room-temperature LB (without antibiotics) to the cells immediately after heat shock.
8. Incubate the cells in a shaking incubator at 30 °C and 225 rpm for 1 h.
9. Plate transformation mixture onto LB agar plates containing 100 μg/mL ampicillin. Spread evenly using a sterile spreader.
10. Incubate the plates overnight at 30 °C.
11. Select the smaller colonies on the plate and culture them in a sterile test tube containing 5 mL of LB media with 100 μg/mL ampicillin.
12. Incubate the culture in a shaking incubator at 30 °C and 225 rpm overnight.
Note: Incubation at 37 °C or choosing larger colonies may lead to plasmids with random DNA insertions and deletions.
B2. CrPV cDNA plasmid purification
1. Harvest 5 mL of E. coli culture grown in LB by centrifuging at 4000× g for 10 min at 4 °C.
2. Extract the plasmid using the NEB Miniprep kit.
3. The plasmid can be validated via a SalI restriction digest and confirming its size by DNA gel electrophoresis.
a. Combine the following components in a sterile 1.5 mL microcentrifuge tube: 10 μg of plasmid DNA, 5 μL of restriction enzyme, 10 μL of 10× reaction buffer, and sterile water to a final volume of 200 μL (adjust volume depending on enzyme and manufacturer’s recommendations).
b. Gently mix the reaction components by pipetting up and down or flicking the tube.
c. Briefly spin the tube to collect the contents at the bottom.
d. Incubate the reaction overnight at 37 °C to allow complete linearization of the plasmid DNA.
e. Cast 1% agarose gel (0.5 g of agarose in 50 mL of 1× TAE buffer) with 1.5 μL of SafeView DNA stain.
f. Add loading dye to 300 ng of samples and load the gel with 1 kb plus DNA ladder.
g. Image the DNA gel using the gel doc to confirm the expected product length.
4. The plasmid should be verified with Sanger sequencing.
B3. CrPV RNA synthesis via in vitro transcription
1. Linearize the CrPV cDNA plasmid by Ecl136II restriction digest, according to the manufacturer’s protocol.
2. Perform DNA clean-up (e.g., Qiagen):
a. In a microfuge tube, add 5× volume of PB buffer to the digested DNA (linearized plasmid).
b. Load the mixture onto a spin column and centrifuge at 13,000× g for 1 min.
c. Wash the column twice with 700 μL of PE wash buffer, spinning at 13,000× g for 1 min after each wash.
d. Elute the cleaned DNA by adding 30–50 μL of DEPC-treated water directly to the column membrane.
e. Centrifuge at 13,000× g for 1 min to collect the eluted DNA.
f. Measure the DNA concentration using a spectrophotometer (e.g., NanoDrop).
3. Verify DNA integrity by running 200–300 ng of DNA on a 1% agarose gel in 1× TAE buffer, as described in step B2.
4. For the in vitro transcription reaction, combine the following components in a sterile 1.5 mL microcentrifuge tube on ice (adjust volumes proportionally for larger reactions): 1 μg of linear DNA, 5 μL of rNTP mix (containing ATP, CTP, GTP, and UTP at 10 mM each), 2 μL of 10× T7 transcription buffer, 0.5 μL of RNase inhibitor (e.g., Ribolock), 0.3 μL of T7 RNA polymerase, and 0.4 μL of YIPP phosphatase.
5. Add DEPC-treated water to bring the final volume to 20 μL.
6. Incubate the reaction at 30 °C for 4 h for RNA synthesis.
7. Perform DNase treatment to remove template DNA:
a. To the completed transcription reaction, add 2.4 μL of DNase buffer and 1 μL of DNase I.
b. Incubate at 37 °C for 1 h to digest the linear DNA template.
8. Confirm RNA production by RNA gel electrophoresis analysis.
a. Prepare 1% agarose gel by adding 0.5 g of agarose to 35 mL of deionized water in a microwave-safe flask or beaker.
i. Heat the mixture until it is almost boiling and the agarose is completely dissolved (using a microwave or hot plate).
ii. Once the agarose solution has cooled slightly (to approximately 60 °C), add 1 μL of Safe-View stain to the gel solution and mix thoroughly by swirling the flask.
iii. Add 5 mL of 10× MOPS buffer to the agarose solution.
iv. In a fume hood, measure and add 10 mL of paraformaldehyde to the solution and mix well.
v. Immediately pour the mixture into the gel mold and insert the comb to create wells.
vi. Allow the gel to solidify at room temperature in the fume hood.
b. Prepare RNA ladder by mixing 1 μL of RiboRuler RNA Ladder with 2 μL of loading dye. For each RNA sample, mix the appropriate volume of RNA (200–300 ng) with loading dye.
c. Heat the RNA samples at 65 °C for 3 min to denature any secondary structures. Increase heating time if secondary structures were observed in previous gels.
d. To load the gel, place the solidified gel in the electrophoresis tank filled with 1× MOPS running buffer.
e. Load the prepared RNA ladder and RNA samples into the wells of the gel.
f. Run the gel at an appropriate voltage (e.g., 100–120 V) until the dye front has migrated sufficiently (typically 1–2 h).
g. Once the run is complete, carefully remove the gel from the tank.
h. Visualize the RNA bands using a gel imaging system (Bio-Rad Gel Doc 2000) to confirm the expected size of the RNA product.
9. Perform RNA purification using a column-based NEB RNA Clean-Up kit.
10. Measure the RNA concentration using a spectrophotometer (e.g., NanoDrop) to determine yield and quality.
B4. CrPV RNA transfection into S2 cells
1. On day 1, using a serological pipette, resuspend Drosophila S2 cells in complete Schneider’s medium in a flask.
2. Count the cells using a hemocytometer or automated cell counter.
3. Seed the S2 cells by plating 2 × 10 cells per well in a 6-well plate.
4. Allow cells to settle by incubating the plate at 25 °C overnight to allow the cells to adhere and settle.
5. On day 2, check under the microscope to ensure that the S2 cells are settled and adherent to the plate.
6. In the biosafety cabinet, prepare transfection solutions.
a. For each transfection, prepare the following solutions in sterile microfuge tubes separately: Solution A (per sample) contains 125 μL of Opti-MEM and 2.5 μL of Lipofectamine 2000. Solution B (per sample) contains 125 μL of Opti-MEM and 3 μg of CrPV RNA.
b. Mix both solutions gently by pipetting up and down.
c. Incubate both solutions separately for 5 min at room temperature.
d. Combine the transfection mixture by adding Solution A to Solution B.
e. Mix gently by pipetting up and down (do not vortex).
f. Incubate the combined solution for 15 min at room temperature to allow complexes to form.
7. After the 15 min incubation, carefully aspirate all of the media from each well containing S2 cells.
8. Pipette to the side of the well to avoid disturbing the cell monolayer during media removal.
9. Gently add the transfection mixture (250 μL) dropwise to each well.
10. Rock the plate gently at room temperature for 15 min to distribute the complexes evenly.
11. Add 2 mL of complete Schneider’s medium (with serum) per well to dilute the transfection mixture.
12. Incubate the plate at 25 °C for 48 h.
13. Observe the S2 cells over time to check for signs of infection, such as membrane blebbing, cell detachment, clumping, or lysis, as described in Kerr et al. [3].
14. When cell lysis is observed, using a serological pipette, detach cells by gently tapping the flask and collect the entire culture in a 15 mL Falcon tube.
15. Centrifuge at 500× g for 5 min to pellet cells, carefully remove the supernatant, and discard it in 10% bleach.
16. Resuspend the cell pellet in 500 μL of sterile PBS.
17. Subject the cell suspension to three freeze-thaw cycles (in a -80 °C freezer followed by a 37 °C water bath) to lyse cells and release virus particles.
18. Centrifuge the lysate at 1,000× g for 10 min at 4 °C to pellet cell debris and collect the supernatant.
19. Aliquot 50 μL of the virus-containing supernatant into PCR strip tubes and store at -80 °C for virus propagation.
C. CrPV propagation in S2 cells
Note: All procedures should be performed in a biosafety cabinet using aseptic techniques.
1. On Day 1, transfer 30 million S2 cells to a 15 mL Falcon tube and centrifuge at 500× g for 5 min.
2. Wash the cell pellet with 5 mL of sterile PBS, centrifuge again, and remove the supernatant.
3. Add the 50 μL virus stock obtained in the previous section to the cell pellet and gently resuspend with a micropipette. Transfer the virus–cell mixture to a microfuge tube.
4. Incubate the mixture at room temperature for 30 min.
5. Optional: During incubation, the microfuge tube can be placed on a rocking platform with gentle rocking (speed at 500× g and angle at 10°) to ensure even distribution of the virus.
6. Transfer the virus–cell mixture to a new T25 flask containing 5 mL of complete S2 cell media.
7. Incubate the flask at 28 °C for 18–24 h to allow virus propagation.
8. On Day 2, detach cells by gently tapping the flask and collect the entire culture in a 15 mL Falcon tube.
9. Centrifuge at 500× g for 5 min to pellet cells, carefully remove the supernatant, and discard it in 10% bleach.
10. Resuspend the cell pellet in 1 mL of sterile PBS.
11. Subject the cell suspension to three freeze-thaw cycles (in a -80 °C freezer followed by a 37 °C water bath) to lyse cells and release virus particles. Option: Freezing the cell suspension in liquid N2 is also appropriate if available.
12. Centrifuge the lysate at 1,000× g for 10 min at 4 °C to pellet cell debris.
13. Collect the virus-containing supernatant and add it to a T75 flask containing S2 cells at 80% confluency.
14. Incubate the flask at 28 °C for 18–24 h to amplify the virus.
15. On Day 3, detach cells and transfer the entire culture to a 50 mL Falcon tube.
16. Centrifuge at 500× g for 5 min to pellet cells, carefully remove the supernatant, and discard it in 10% bleach.
17. Resuspend the cell pellet in 2 mL of sterile PBS.
18. Perform three freeze-thaw cycles as described previously to release virus particles.
19. Centrifuge the lysate at 1,000× g for 10 min at 4 °C and collect the supernatant.
20. Aliquot 10–30 μL of the virus-containing supernatant into PCR strip tubes and store at -80 °C.
D. Confirmation of CrPV genome via Sanger sequencing
1. In the chemical fume hood, add 500 μL of Trizol to 20 μL of virus stock. Transfer all volume into a 1.5 mL microfuge tube.
2. In the fume hood, add 200 μL of chloroform to the sample. Mix well by vortexing and incubate at room temperature for 2 min.
3. Centrifuge at 13,000× g for 15 min at 4 °C.
4. Transfer 400 μL of the top aqueous layer to a new 1.5 mL microfuge tube (around 80% of the top layer).
5. Add 400 μL of isopropanol to the sample. Mix well by vortexing and incubate at room temperature for 10 min.
6. Spin down the RNA at 13,000× g for 30 min at 4 °C (a small white RNA pellet may be visualized).
7. Remove the supernatant without disturbing the pellet.
8. Rinse the pellet by pipetting up and down with >200 μL of 75% ethanol and then centrifuge at 13,000× g for 5 min.
9. Air dry the pellet for 2 min (do not completely dry).
10. Resuspend in 20 μL of RNase-free water.
11. Measure the concentration of the RNA using the RNA Qubit Quantification kit.
12. Reverse transcribe 500 ng of viral RNA to cDNA using the LunaScript RT SuperMix kit.
13. Purify cDNA using the Monarch PCR & DNA Cleanup kit.
14. Measure cDNA concentration using a nanodrop.
15. Using primers specifically covering the region of interest (Figure 2A), perform PCR using the Phusion High-Fidelity PCR kit. Primers, CrPV-Seq2, and CrPV-Seq26R are used in this demonstration (Figure 2B).
Figure 2. PCR amplification of cricket paralysis virus CrPV-1A region for wildtype (WT) and a mutant CrPV-1A (R146A). A. Schematic representation of the CrPV genome, highlighting the positions of primers used for amplifying specific regions. CrPV-Seq1 to CrPV-Seq28R target various segments of the CrPV genome, ensuring complete coverage for sequencing and validation. B. PCR product analysis for the CrPV-1A region from WT and R146A virus. The CrPV-1A region, which is critical for viral replication, is amplified in both the WT CrPV and the R146A mutant. Gel electrophoresis shows the successful amplification of a ~1 kb product from both WT CrPV and the R146A mutant, using CrPV-Seq2 and CrPV-Seq26R, confirming that the mutation does not interfere with the amplification of the targeted region. This PCR product is used for further validation via Sanger sequencing.
16. Purify the PCR product using the Monarch PCR & DNA Cleanup kit.
17. Measure the amplicon concentration using a nanodrop.
18. Run DNA gel electrophoresis to validate the size of the PCR product (Figure 2B).
a. Cast 1% agarose gel (0.5 g of agarose in 50 mL of 1× TAE buffer) with 1.5 μL of SafeView DNA stain.
b. Add loading dye to 300 ng of PCR samples and load the gel with either 1 kb or 100 bp DNA ladder based on the size of the targeted region.
c. Image the DNA gel using the gel doc to confirm the expected product length.
19. Verify the sequence through Sanger sequencing.
20. Align sequencing results to CrPV genome on SnapGene.
E. Determination of CrPV viral titer via immunofluorescence-based focus-forming unit (FFU) assay
1. Prepare serial dilutions of the viral stock from 100 to 10-8 in 1× PBS (90 μL of PBS, 10 μL of virus).
2. Coat a 96-well plate with 50 μL of ConA (0.5 mg/mL in PBS, filter-sterilized) per well and incubate for at least 60 min at room temperature in the biological safety cabinet.
3. Aspirate the ConA solution and allow the plate to air dry.
4. Prepare microfuge tubes containing 1.8 × 106 S2 cells each and pellet cells by centrifugation (500× g for 30 s).
5. Add 10 μL of each virus dilution (or PBS for mock) to the cell pellets and mix gently to resuspend.
6. Incubate the virus–cell mixtures for 30 min at 25 °C and then add 600 μL of media to each tube.
7. Following the map in Figure 3, transfer 100 μL of infected cells to each well of the ConA-coated plate and incubate for 8 h at 28 °C (plate 300,000 cells per well).
8. Aspirate the media and wash the cells with 1× PBS.
9. Fix cells with 50 μL of 3% paraformaldehyde in FBS for 15 min at room temperature, then wash with 100 μL of 1× PBS. The plates can be stored at 4 °C for up to a week.
10. Permeabilize cells with 50 μL of methanol for 10 min at room temperature, then wash with 100 μL of 1× PBS.
11. Incubate cells with 50 μL of primary antibody against ORF2 (anti-VP2, 1:500 in 5% BSA/PBS) for 1 h at room temperature.
12. Wash cells three times with 100 μL of 1× PBS.
13. Incubate cells with 50 μL of Texas red goat anti-rabbit secondary antibody (1:500 in 5% BSA/PBS) for 1 h at room temperature in the dark.
14. Wash cells twice with 100 μL of 1× PBS.
15. Stain nuclei with 50 μL of Hoechst dye (1:20,000 in PBS) for 15 min at room temperature.
16. Wash once with 100 μL of 1× PBS and store the plate at 4 °C in 100 μL of 1× PBS, protected from light by covering the plate with aluminum foil.
17. For storage, fill the outer circle of wells labeled with the blue box (Figure 3) with 100 μL of water in each well to prevent sample evaporation.
Figure 3. Serial dilution of cricket paralysis virus (CrPV) for focus-forming assay (FFA). A 96-well plate layout illustrating the serial dilutions of CrPV used for the focus-forming assay (FFA). The virus stock is serially diluted from 100 to 10-8 across columns 1 to 9, with each dilution plated in quadruplicate. Column 10 contains mock-infected wells (no virus control), used to ensure the absence of contamination and background signal. This setup allows for the quantification of CrPV titers by counting the number of focus-forming units (FFU) in each well after staining with Texas red-conjugated antibodies against CrPV capsid protein (VP2) and performing immunofluorescence imaging.
F. Imaging and cell counting via fluorescence microscopy (i.e., CX5 HCS platform)
Note: Any high-quality fluorescence microscopy, such as an automated fluorescence microscope or standard fluorescence microscope equipped with appropriate filters, can be used. At the UBC Life Sciences Institute, we have access to a CellInsight CX5 HCS platform for imaging and cell counting. As such, we describe procedures specific to this equipment.
1. Prior to imaging, confirm immunofluorescence on an EVOS microscopic imaging system or equivalent (Figure 4A).
Figure 4. Immunofluorescence detection of cricket paralysis virus (CrPV) capsid protein (VP2) in infected S2 cells. A. Immunofluorescence EVOS images of S2 cells infected with wildtype (WT) CrPV at a dilution of 10-1, compared to mock-infected control (no CrPV infection), with the scale bar set to 150 μm. The CrPV VP2 capsid protein is detected using Texas red-conjugated antibodies, and the cell nuclei are stained with Hoechst dye (blue). WT CrPV-infected cells show significant VP2 expression (red), while the mock-infected cells show no VP2 signal. B. Immunofluorescence CX5 images of S2 cells infected with WT CrPV at two different dilutions, 10-1 and 10-4, compared to mock-infected cells, with the scale bar set to 100 μm. The Hoechst-stained nuclei (blue) are consistent across all conditions. CrPV VP2 (Texas red) expression increases with higher viral loads, as seen in the 10-1 infected cells, whereas the mock-infected cells show no capsid protein expression. The figure demonstrates the progression of CrPV infection and viral protein expression as a function of viral dilution.
2. Acquire images on the CellInsignt CX5 HCS Platform. The following steps are specific to the 2015 version of the HCS Navigator system.
3. On the CellInsight CX5 HCS Platform, power on the HCS Navigator system and ensure proper connection to the microscope and camera.
4. Launch HCS Navigator and select the scan plate on the connected computer.
5. Choose Fluorescence as the imaging mode and select 96-well plate format as the plate layout.
6. To configure fluorescence channels, keep channel 1 configured for white light illumination to help identify areas with an optimal density of cells for imaging. For channel 2 configuration, select an excitation of 350 nm and emission around 461 nm for Hoechst. For channel 3 configuration, select an excitation of 595 nm and emission around 615 nm for Texas red.
7. Adjust the exposure times for each channel to optimize the signal-to-noise ratio.
8. Perform a test capture on a sample well to ensure correct exposure for both the Texas red and Hoechst channels. Sample images are shown in Figure 4B.
9. Maintain a consistent exposure time throughout the entire imaging process.
10. Use the software’s auto-focus function to focus on the cells in each of the wells.
11. Create a scan plan to capture three fields per well and define four wells per dilution for imaging (this yields a technical replicate of three in four biological replicates, collecting 12 images for each virus sample).
12. Begin the imaging run by starting the scan plan.
13. Monitor the initial images to ensure proper focus and exposure.
14. Review the acquired images for any anomalies or focus issues.
15. If needed, select different fields and re-image specific wells.
16. Open the captured images from the Hoechst channel in the analysis platform.
17. Select the cell counting tool and set counting parameters by selecting the radius of the size of the nuclei.
18. Execute the cell counting analysis on the entire dataset to obtain the total number of cells per image field.
19. Save the image data and metadata to the designated storage location.
20. Export images as TIFF and record data from the analysis for downstream titer calculation.
Note: Selecting a dilution of 10-5 or less ensures that the virus concentration is within a manageable range for detection, counting, and accurate determination. Using a dilution that is too low (high virus concentration) can cause overcrowding, multiple infections per cell, signal saturation, inaccurate titer estimation, and difficulty determining the endpoint of infection (Figure 4B).
G. Viral titer calculation
1. Open the Texas red images from CX5 in ImageJ and adjust the brightness and contrast to count the number of “red cells” in the image. The red cells are the ones that are CrPV-positive.
2. Record the number of positive cells.
3. Follow the sample calculation on the attached viral titer calculation Excel sheet.
4. Calculate the percentage of positive cells by dividing the red cell count (infected cells stained with Texas red) by the total cell count (stained with Hoechst). Keep this value as a decimal.
Formula: = Viral-infected Cell#/Total Cell#
5. Determine the total number of positive cells in the well by multiplying the percentage of positive cells by the total number of cells plated in the well (e.g., 300,000 cells).
Formula: =Percentage_Positive_Cells * 300000
6. Adjust for infection volume by multiplying the total number of positive cells by the inverse of the infection volume (e.g., for 10 μL infection volume, multiply by 1/10).
Formula: =Total_Positive_Cells * (1/Infection_Volume)
7. Account for the dilution factor by multiplying the adjusted positive cell count by the dilution factor used during the experiment.
Formula: =Adjusted_Positive_Cells * Dilution_Factor
8. Convert the final value to scientific notation to obtain the viral titer in FFU/μL (focus forming units per microliter).
H. Validation of CrPV infection by RNA gel electrophoresis and western blotting
H1. CrPV infection
1. Using the calculated viral titer, infect S2 cells at MOI = 10 for the desired amount of time.
2. Prepare microfuge tubes containing 3 × 106 S2 cells each and pellet cells by centrifugation (500× g for 30 s).
3. Add the calculated volume of virus to the cell pellets and mix gently to resuspend.
4. Incubate the virus–cell mixtures for 30 min at 25 °C for absorption; then, add 500 μL of media to each tube.
5. Split the total volume of infected cells equally to two wells of a 6-well plate (2 mL media per well) and incubate at 25 °C for the desired amount of time.
6. Monitor the cytopathic effect under a microscope (Figure 5A).
7. Transfer all cells from those two wells, combining them into a 15 mL tube, and pellet cells by centrifugation (500× g for 3 min).
8. Aspirate media and wash cells with 5 mL of 1× PBS buffer.
9. Pellet cells by centrifugation in a tabletop centrifuge (e.g., microfuge) (500× g for 3 min) and aspirate the PBS.
H2. Sample preparation (whole-cell RNA extraction)
1. Follow the Trizol RNA isolation procedure as described in Section D.
H3. Sample preparation (whole-cell protein extraction)
1. Mix the cell pellet in 50 μL of RIPA buffer containing 1× protease inhibitor cocktail (50 μL of RIPA buffer per 5 million cells).
2. Store samples in Eppendorf tubes at -80 °C until ready to proceed.
3. Freeze-thaw samples three times at -80 °C and 37 °C to fully lyse cells.
4. Centrifuge at 13,000× g for 30 min at 4 °C to pellet cell debris.
5. Transfer the aqueous layer to the new tubes, being careful to avoid the cell pellet (it will be sticky and goopy).
6. Proceed with the Bradford assay to measure protein concentration.
a. Prepare a BSA standard curve with concentrations of 0, 2, 4, 6, 8, and 10 μg/μL in distilled water.
b. Make a 1:5 dilution of protein assay dye in water.
c. Prepare cuvettes with 1 mL of diluted dye (1 cuvette per sample, plus 1 for each BSA standard and 1 for the blank).
d. Add 2–5 μL of BSA standards or protein lysate to each cuvette and mix by pipetting with a P1000.
e. Measure the OD at 595 nm using the spectrophotometer set to Bradford Assay.
f. Plot the BSA standard curve to determine the relationship between OD and protein concentration.
g. Use the standard curve to determine the protein concentration of the samples and calculate the amount needed for 10 μg of protein.
7. Add 2× blue protein loading dye to the protein lysate and mix well.
8. Boil samples at 95 °C for 5 min.
9. Samples are ready for loading on an SDS-PAGE gel for western blotting or can be stored in the -20 °C freezer until ready to proceed.
H4. RNA gel electrophoresis
1. Prepare 1% agarose gel as described in the previous section.
2. Prepare the RNA ladder by mixing 1 μL of RiboRuler RNA ladder with 2 μL of loading dye. For each RNA sample, mix the appropriate volume of RNA (200–300 ng) with loading dye.
3. Heat the RNA samples at 65 °C for 3 min to denature any secondary structures. Increase heating time if secondary structures were observed in previous gels.
4. To load the gel, place the solidified gel in the electrophoresis tank filled with 1× MOPS running buffer.
5. Load the prepared RNA ladder and RNA samples into the wells of the gel.
6. Run the gel at an appropriate voltage (e.g., 100–120 V) until the dye front has migrated sufficiently (typically 1–2 h).
7. Once the run is complete, carefully remove the gel from the tank.
8. Visualize the RNA bands using a gel imaging system (Bio-Rad Gel Doc 2000). Use appropriate filters/settings for Safe-View stain (Figure 5B). Figure 5B shows an example RNA gel electrophoresis image of total RNA extracted from S2 cells infected with wildtype (WT) CrPV and the CrPV mutant R146A at different time points. Clear bands of CrPV RNA are visible in both WT and R146A-infected cells, confirming active viral replication. The mock-infected sample serves as a control, displaying only S2 ribosomal RNA and transfer RNA bands, without any CrPV RNA, indicating no infection.
Figure 5. Infection and RNA analysis of cricket paralysis virus (CrPV)-infected and mutant CrPV-1A (R146A)-infected S2 cells. A. EVOS brightfield images of S2 cells showing the progression of CrPV infection, with scale bars set to 150 μm. Uninfected S2 cells are shown as a control. Cells infected with CrPV at a multiplicity of infection (MOI) of 10 for 9 h show early cytopathic effects, while cells infected for 15 h display more pronounced cell death and morphological changes due to viral infection. B. RNA gel electrophoresis of total RNA extracted from S2 cells infected with WT CrPV and CrPV 1A (R146A) mutant at MOI of 10, at different time points (3, 6, and 9 h post-infection, h.p.i.). The gel shows clear CrPV RNA bands in both WT CrPV and mutant CrPV (R146A)-infected cells, confirming active viral replication. The mock-infected sample serves as a negative control, showing only S2 ribosomal RNA (rRNA) and transfer RNA (tRNA) bands. The absence of viral RNA bands in the mock sample indicates no viral infection. The CrPV RNA bands demonstrate similar levels of viral replication in both WT CrPV and CrPV (R146A) mutants over time.
H5. Western blotting
1. In separate 50 mL Falcon tubes, prepare the resolving (12%) and stacking (4%) gel solutions.
2. Assemble the gel plates in the casting frame and ensure they are tightly sealed.
3. Place the frame vertically into the gel casting stand.
4. Mix the prepared resolving gel solution by adding APS and TEMED last to initiate polymerization.
5. Pour the resolving gel solution into the gap between the gel plates, leaving about 1.5 cm of space at the top for the stacking gel.
6. Gently overlay the gel with isopropanol to ensure a flat surface and prevent air bubbles.
7. Allow the gel to polymerize for 20–30 min at room temperature.
8. After the resolving gel has polymerized, carefully remove the isopropanol by tilting the gel plates and drying the surface with filter paper.
9. Mix the stacking gel solution by adding APS and TEMED last.
10. Pour the stacking gel on top of the polymerized resolving gel.
11. Insert the gel comb gently into the stacking gel to create wells.
12. Allow the stacking gel to polymerize at room temperature for 15–20 min.
13. Once the stacking gel has polymerized, carefully remove the comb by pulling it straight up.
14. Assemble the gel cassette and fill both the inner and outer chambers with 1× SDS running buffer.
15. Rinse the wells with the running buffer to remove any unpolymerized acrylamide.
16. The gel is now ready to load protein samples and perform electrophoresis.
17. Mix 10–20 μg of protein sample with protein loading dye and heat at 95 °C for 5 min to denature proteins.
18. Load the protein samples and a protein ladder into the wells of the gel.
19. Run the gel at 60 mA until the dye front reaches the bottom (approximately 50 min).
20. Once completed, remove the gel and wash with distilled water to remove the excess buffer.
21. Prepare 1× transfer buffer (with 15% methanol) and soak the PVDF membrane in methanol for 30 s before transferring to cold transfer buffer.
22. Assemble the transfer stack by the order of a sponge, two sheets of Whatman paper, gel (gently remove bubbles), PVDF membrane (gently remove bubbles), two sheets of Whatman paper, and a sponge.
23. Transfer proteins at 100 V for 1.5 h in the cold room.
24. After the transfer, block the membrane in 0.1% TBST with 5% milk for 1 h at room temperature to prevent non-specific binding.
25. Incubate the membrane with the primary antibody specific to CrPV-VP2 (1:500 dilution in 5% milk-TBST) or CrPV-1A (1:1,000 in 5% milk-TBST) or α-Tubulin (1:1,000 in 5% milk-TBST) for 1 h at room temperature or overnight at 4 °C.
26. Wash the membrane three times with 0.1% TBST for 10 min each.
27. Incubate the membrane with the secondary antibody (1:5,000 dilution in 5% milk-TBST) for 1 h.
28. Wash the membrane three times with 0.1% TBST for 5 min each.
29. Pour out the 0.1% TBST, dab the side of the membrane with a Kimwipe to draw off the remaining TBST, then lay flat back in the empty dish.
30. Add ECL (from the chemiluminescence kit) mix on the membrane. Move left-to-right, front-to-back a couple of times to spread it out. Let it sit for 3–4 min at room temperature.
31. Follow the manufacturer's protocol to detect proteins by enhanced chemiluminescence (Figure 6).
Figure 6. Immunoblots of S2 cell lysates infected with cricket paralysis virus (CrPV) and mutant CrPV (R146A). Western blot analysis of lysates from S2 cells infected with either mock (M), wildtype CrPV, or CrPV mutant (R146A) at a multiplicity of infection (MOI) of 10, collected at indicated time points. The detection of viral protein expression in the infected cell lysates confirmed that the propagated virus is infectious. This figure is adapted from Sadasivan et al. [5].
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
• Sadasivan et al. [5]. Targeting Nup358/RanBP2 by a viral protein disrupts stress granule formation. PLoS Pathogens.
• Warsaba et al. [17]. Multiple Viral Protein Genome-Linked Proteins Compensate for Viral Translation in a Positive-Sense Single-Stranded RNA Virus Infection. Journal of Virology.
• Kirby et al. [9]. The Hinge Region of the Israeli Acute Paralysis Virus Internal Ribosome Entry Site Directs Ribosomal Positioning, Translational Activity, and Virus Infection. Journal of Virology.
• Wang et al. [10]. Resurrection of a viral internal ribosome entry site from a 700-year-old ancient Northwest Territories Cripavirus. Viruses.
• Kerr et al. [11]. Transmission of Cricket paralysis virus via exosome-like vesicles during infection of Drosophila cells. Scientific Reports.
• Kerr et al. [12]. IRES-dependent ribosome repositioning directs translation of a +1 overlapping ORF that enhances viral infection. Nucleic Acids Research.
• Nayak et al. [13]. A viral protein restricts Drosophila RNAi immunity by regulating argonaute activity and stability. Cell Host & Microbe.
• Au et al. [14]. Functional insights into the adjacent stem-loop in honey bee dicistroviruses that promotes IRES-mediated translation and viral infection. Journal of Virology.
• Khong et al. [4]. Disruption of stress granule formation by the multifunctional cricket paralysis virus 1A protein. Journal of Virology.
• Kerr et al. [15]. Molecular analysis of the factorless internal ribosome entry site in Cricket Paralysis virus infection. Scientific Reports.
• Khong et al. [16]. Temporal regulation of distinct internal ribosome entry sites of the dicistrovirus cricket paralysis virus. Viruses.
• Kerr et al. [3]. The 5' untranslated region of a novel infectious molecular clone of the dicistrovirus cricket paralysis virus modulates infection. Journal of Virology.
General notes and troubleshooting
1. Work with cricket paralysis virus (CrPV) in our lab is performed under Biosafety Level 2 (BSL-2) conditions, as CrPV is not pathogenic to humans. However, safety regulations may vary among countries and institutions. Researchers intending to work with this virus should consult their local biosafety office to determine the appropriate safety requirements before starting any experiments.
2. For S2 cell maintenance, avoid splitting cells below a density of 0.5 × 10 cells/mL, as under-confluent cells may take an extended time to recover or fail to recover entirely. Passaging cells regularly (every 2–4 days) maintains them in the logarithmic growth phase and ensures a doubling time of approximately 24 h.
3. To increase the yield during CrPV cDNA plasmid extraction using the NEB Miniprep Kit, preheat the elution buffer to 50 °C before use.
4. To acquire fluorescence images using the CellInsight CX5 HCS platform, general instructions are available through Thermo Fisher Scientific support or built-in help features within the HCS Navigator software. Additionally, instructions can be found on the Thermo Fisher Scientific website:
https://www.thermofisher.com/ca/en/home/life-science/cell-analysis/cellular-imaging/high-content-screening/cellinsight-cx5.html.
We are using the 2015 version of the system, which is no longer available for installation. Therefore, we recommend that users refer to online resources or contact the manufacturer for the most up-to-date instructions and support.
5. If immunofluorescence images of mock-infected CX5 samples show high background noise in the CrPV-VP2 channel, ensure that S2 cells are at approximately 70%–80% confluency before infection. If the issue persists, perform additional wash steps using PBS buffer after antibody incubation, and carefully pipette from the side of the wells in the 96-well plate to avoid disturbing the cell monolayer.
Acknowledgments
This study was supported by the Canadian Institutes of Health Research (PJT-178342 to E.J.), the National Sciences and Engineering Research Council of Canada (RGPIN-2023-03658 to E.J.) and the SERB-UBC PhD award to J.S.
We acknowledge the authors of the previously published work from which Figure 2 is adapted, Warsaba et al. [2], and the authors of the previously published work from which Figure 7 is adapted, Sadasivan et al. [5].
Competing interests
Authors declare no competing interests.
References
Bonning, B. C. and Miller, W. A. (2010). Dicistroviruses. Annu Rev Entomol. 55(1): 129–150.
Warsaba, R., Sadasivan, J. and Jan, E. (2020). Dicistrovirus-Host Molecular Interactions. Curr Issues Mol Biol. 34: 83–112.
Kerr, C. H., Wang, Q. S., Keatings, K., Khong, A., Allan, D., Yip, C. K., Foster, L. J. and Jan, E. (2015). The 5′ Untranslated Region of a Novel Infectious Molecular Clone of the Dicistrovirus Cricket Paralysis Virus Modulates Infection. J Virol. 89(11): 5919–5934.
Khong, A., Kerr, C. H., Yeung, C. H. L., Keatings, K., Nayak, A., Allan, D. W. and Jan, E. (2017). Disruption of Stress Granule Formation by the Multifunctional Cricket Paralysis Virus 1A Protein. J Virol. 91(5): e01779–16.
Sadasivan, J., Vlok, M., Wang, X., Nayak, A., Andino, R. and Jan, E. (2022). Targeting Nup358/RanBP2 by a viral protein disrupts stress granule formation. PLoS Pathog. 18(12): e1010598.
Sadasivan, J., Hyrina, A., DaSilva, R. and Jan, E. (2023). An Insect Viral Protein Disrupts Stress Granule Formation in Mammalian Cells. J Mol Biol. 435(16): 168042.
Nonis, S. G., Haywood, J., Schmidberger, J. W., Mackie, E. R. R., Soares da Costa, T. P., Bond, C. S. and Mylne, J. S. (2021). Structural and biochemical analyses of concanavalin A circular permutation by jack bean asparaginyl endopeptidase. Plant Cell. 33(8): 2794–2811.
Garrey, J. L., Lee, Y. Y., Au, H. H. T., Bushell, M. and Jan, E. (2010). Host and Viral Translational Mechanisms during Cricket Paralysis Virus Infection. J Virol. 84(2): 1124–1138.
Kirby, M. P., Stevenson, C., Worrall, L. J., Chen, Y., Young, C., Youm, J., Strynadka, N. C. J., Allan, D. W. and Jan, E. (2022). The Hinge Region of the Israeli Acute Paralysis Virus Internal Ribosome Entry Site Directs Ribosomal Positioning, Translational Activity, and Virus Infection. J Virol. 96(5): e01330–21.
Wang, X. and Jan, E. (2021). Resurrection of a viral internal ribosome entry site from a 700-year-old ancient Northwest Territories cripavirus. Viruses. e428736.
Kerr, C. H., Dalwadi, U., Scott, N. E., Yip, C. K., Foster, L. J. and Jan, E. (2018). Transmission of Cricket paralysis virus via exosome-like vesicles during infection of Drosophila cells. Sci Rep. 8(1): 17353.
Kerr, C. H., Wang, Q. S., Moon, K. M., Keatings, K., Allan, D. W., Foster, L. J. and Jan, E. (2018). IRES-dependent ribosome repositioning directs translation of a +1 overlapping ORF that enhances viral infection. Nucleic Acids Res. 46(22): 11952–11967.
Nayak et al. (2018). A Viral Protein Restricts Drosophila RNAi Immunity by Regulating Argonaute Activity and Stability. Cell Host Microbe. 24(4): 542–557.e9.
Au, H. H. T., Elspass, V. M. and Jan, E. (2018). Functional Insights into the Adjacent Stem-Loop in Honey Bee Dicistroviruses That Promotes Internal Ribosome Entry Site-Mediated Translation and Viral Infection. J Virol. 92(2): e01725–17.
Kerr, C. H., Ma, Z. W., Jang, C. J., Thompson, S. R. and Jan, E. (2016). Molecular analysis of the factorless internal ribosome entry site in Cricket Paralysis virus infection. Sci Rep. 6(1): e1038/srep37319.
Khong, A., Bonderoff, J., Spriggs, R., Tammpere, E., Kerr, C., Jackson, T., Willis, A. and Jan, E. (2016). Temporal Regulation of Distinct Internal Ribosome Entry Sites of the Dicistroviridae Cricket Paralysis Virus. Viruses. 8(1): 25.
Warsaba, R., Stoynov, N., Moon, K. M., Flibotte, S., Foster, L. and Jan, E. (2022). Multiple Viral Protein Genome-Linked Proteins Compensate for Viral Translation in a Positive-Sense Single-Stranded RNA Virus Infection. J Virol. 96(17): e00699–22.
Article Information
Publication history
Received: Nov 1, 2024
Accepted: Jan 8, 2025
Available online: Jan 22, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Correction Notice: A Microplate-Based Expression Monitoring System for Arabidopsis NITRATE TRANSPORTER2.1 Using the Luciferase Reporter
YU Yoshiaki Ueda
SY Shuichi Yanagisawa
Published: Jan 20, 2025
DOI: 10.21769/BioProtoc.5217 Views: 97
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After official publication in Bio-protocol (https://bio-protocol.org/e5127), the following correction has been made:
In Figure 2, the label “Relative fluorescence units” has been corrected to “Relative luminescence units”.
In section “Data analysis”, the term “fluorescence units” has been corrected to “luminescence units”.
Article Information
Publication history
Published: Jan 20, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Streamlined Quantification of Microglial Morphology in Mouse Brains Using 3D Immunofluorescence Analysis
MD Maria Helena de Donato
AK Armin Kouchaeknejad
AD Andreu de Donato
GW Gunter Van Der Walt
EP Emma Puighermanal
In Press, Available online: Jan 26, 2025
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Abstract
Microglial cells are crucial patrolling immune cells in the brain and pivotal contributors to neuroinflammation during pathogenic or degenerative stress. Microglia exhibit a heterogeneous "dendrite-like" dense morphology that is subject to change depending on inflammatory status. Understanding the association between microglial morphology, reactivity, and neuropathology is key to informing treatment design in diverse neurodegenerative conditions from inherited encephalopathies to traumatic brain injuries. However, existing protocols for microglial morphology analyses lack standardization and are too complex and time-consuming for widescale adoption. Here, we describe a customized pipeline to quantitatively assess intricate microglial architecture in three dimensions under various conditions. This user-friendly workflow, comprising standard immunofluorescence staining, built-in functions of standard microscopy image analysis software, and custom Python scripts for data analysis, allows the measurement of important morphological parameters such as soma and dendrite volumes and branching levels for users of all skill levels. Overall, this protocol aims to simplify the quantification of the continuum of microglial pathogenic morphologies in biological and pharmacological studies, toward standardization of microglial morphometrics and improved inter-study comparability.
Key features
• Comparison of 3D microglial architecture between physiological and pathological conditions.
• Quantitative assessment of critical microglial morphological features, including soma volume, dendrite volume, branch level, and filament length.
• Simplified, semi-automated data export and analysis through simple Python scripts.
Keywords: Microglia Imaris Morphology Neuroinflammation Cell phenotype Brain
Graphical overview
Background
Neuroinflammation, a main mechanism behind neuronal death and dysfunction in various pathologies, involves microglia as key players patrolling immune cells [1]. These cells, known for their morphological heterogeneity and dynamic processes, play diverse roles in different neurodegenerative disorders [2,3]. It is well-known that the functional and morphological alterations in response to intracellular or extracellular cues are highly interconnected. For example, microglia often exhibit increased volume and altered dendrite architecture when performing phagocytic functions in damaged loci, as opposed to a ramified, homeostatic morphology in healthy brain tissue [4,5].
Given the fundamental role of microglia as resident immune cells in the central nervous system, understanding their behavior in pathological contexts is crucial. In this regard, light microscopy–based immunohistochemical (IHC) analyses of ex vivo brain sections remain a core tool for neuropathology research centers worldwide. However, the dense and intricate microglial architecture poses challenges for 3D morphological studies, necessitating a workflow that builds on standard techniques and tools to facilitate standardization and widespread adoption in neuropathology research. Previous methods for 3D IHC analyses have inherent limitations that result in high error rates when measuring numerous small parameters in large and complex brain samples. This often requires significant time investments, specialized analyses, and multiple software tools. Moreover, commonly used techniques for quantitative microglial assessment lack data standardization, resulting in non-comparable results [5]. This is exemplified by the inherent limitations of common open-source tools, such as ImageJ, which either lack the capability for 3D assessment or require complex pipelines involving non-standardized plug-ins to obtain 3D data [3,6–9].
To overcome these challenges and capture the continuum of microglial morphologies under various physiological and pathological states, we describe a straightforward and repeatable custom approach for 3D microglial morphology analyses in mouse brain sections. Our pipeline is based on 3D image acquisition workflows built into the standard Imaris software, followed by custom Python scripts for data analysis. Similar approaches have been successfully employed for 3D analyses of cell volumes and morphological features, albeit in distinct research fields from neuroscience [10,11]. The protocol described here was used to study neuroinflammation and treatment response in primary mitochondrial disorders, specifically in a mouse model of Leigh syndrome that presents distinct patterns of inflammatory neurodegeneration associated with treatment-resistant epilepsy [12,13].
This workflow may be applicable to multiple conditions, providing biological plausibility for the association between gene disruptions and neuropathological phenotypes. It also enables the assessment of the effects of various pharmacological treatments, thereby informing pre-clinical drug development.
Materials and reagents
Biological materials
1. Mice with conditional Ndusf4 ablation in Gad2-espressing neurons (Gad2Cre/+:Ndufs4lox/lox or Gad2:Ndufs4cKO) were obtained as previously described [12,13]. Gad2Cre/+:Ndufs4lox/+ healthy males, expressing Cre specifically in Gad2-positive cells, were crossed with healthy females that have floxed Ndufs4 exon 2 (Gad2+/+, Ndufs4lox/lox), resulting in Gad2:Ndufs4cKO pups or sex/age-matched controls lacking Cre-expression (Gad2+/+) or functional loxP pair on the target gene (Ndufs4lox/+). A full description of the analyses and results pertaining to these samples is described in [13].
Reagents
1. Ketamine (UAB, catalog number: 579557)
2. Sodium chloride (NaCl) (Acros Organics, catalog number: 207790010)
3. Xylazine (Bayer, catalog number: 572126)
4. Phosphate buffered saline (PBS) (Millipore, catalog number: 524650)
5. Paraformaldehyde (PFA) (Sigma-Aldrich, catalog number: 441244)
6. Ethylene glycol (Merck, catalog number: 102466)
7. Glycerol (Fisher Scientific, catalog number: 12144481)
8. Sodium phosphate (NaH2PO4) (Merck, catalog number: S5011)
9. Triton X-100 (Sigma-Aldrich, catalog number: X100)
10. Bovine serum albumin (BSA) (Sigma-Aldrich, catalog number: 810533)
11. DAPI-Fluoromount-G solution (Electron Microscopy Science, catalog number: 17984-24)
12. Primary antibody, IBA1 (Wako, catalog number: 019-19741)
13. Secondary antibody, Alexa Fluor 647 (Fisher Scientific, catalog number: A-31573)
Solutions
1. Anesthesia solution (see Recipes)
2. Anti-freezing solution (see Recipes)
3. Permeabilization solution (see Recipes)
4. Blocking solution (see Recipes)
5. Antibody solution (see Recipes)
Recipes
1. Anesthesia solution
100 mg/mL ketamine
20 mg/mL xylazine
0.9% NaCl
2. Anti-freezing solution
30% ethylene glycol
30% glycerol
0.1 M sodium phosphate
3. Permeabilization solution
0.2% Triton X-100
0.1 M PBS
4. Blocking solution
3% BSA
0.1 M PBS
5. Antibody solution
1% BSA
0.15% Triton X-100
0.1 M PBS
Laboratory supplies
1. 24-well plates (Fisher Scientific, catalog number: 11835275)
2. Slides (Fisher Scientific, catalog number: J1800AMNZ)
3. Cover glasses (VWR, catalog number: 631-1339)
Equipment
1. Confocal microscope (Zeiss, model: LSM780)
2. Peristaltic pump for mouse perfusion (Gilson Inc, model: Gilson Minipuls 2)
3. Vibratome (Leica, model: VT 1000S Vibrating-blade microtome)
Software and datasets
1. Imaris software (Bitplane, Belfast, UK, v.9.5, October 17, 2019) with modules for data management, visualization, 3D rendering, and analysis
2. Zeiss LSM Image Browser (version LSM780, 2010)
a. Python 3.11.2 (February 8, 2023)
b. Microsoft Excel (Microsoft, Redmon, WA, Office 365, April 2023)
Code and software availability
Imaris software can be obtained with a license key from the manufacturer (BitPlane, under Oxford Instruments, Belfast, UK). The Python scripts developed here are available from GitHub: https://github.com/AndreudeDonato/Read_Imaris_zstack.git
Data availability
All data shown here are from Puighermanal et al. [13] and are available online at https://doi.org/10.1038/s41467-024-51884-8.
Procedure
A. Tissue preparation for immunofluorescence
1. Anesthesia and perfusion
Anesthetize (intraperitoneally) mice with anesthesia solution (see Recipes) and perfuse (transcardially) with 4% (w/v) PFA in PBS (0.1 M, pH 7.5).
2. Post-fixation
Following perfusion, carefully remove brains, post-fix overnight in 4% PFA solution, and store at 4 °C.
3. Sectioning
After post-fixation, wash brains in PBS and section them into 30 µm-thick sections using a vibratome (Leica). Sections at the level of the external globus pallidus are identified based on anatomical landmarks using a mouse brain atlas, within a window between -0.1 and -0.94 mm from bregma [14].
4. Storage
Store brain sections at -20 °C in anti-freezing solution (see Recipes). This cryoprotectant solution preserves tissue integrity until further processing for immunofluorescence.
B. Immunofluorescence analysis
On day 1, rinse free-floating slices three times in 0.1 M PBS. After 15 min incubation in permeabilization solution (see Recipes), block slices in blocking solution (see Recipes) for 1 h. After the blocking, incubate slices in antibody solution (see Recipes) for 24 h at 4 °C with the microglia/macrophage marker IBA1 (1:1,000).
On day 2, rinse sections three times for 10 min in PBS and incubate for 45–60 min with an Alexa Fluor 647 secondary antibody (1:750). Rinse sections for 10 min twice in PBS and twice in phosphate buffer (0.1 M, pH 7.5) before mounting in DAPI-Fluoromount-G solution [16].
C. Image acquisition
Capture images of IBA1 immunofluorescence with a LSM780 confocal microscope as described [15] using the Z-stack functionality over a span of ~20 μm with intervals of 0.8 μm. The chosen interval of 0.8 μm was selected to ensure no loss of information and to achieve accurate 3D reconstruction of microglial morphology. This resolution is critical for maintaining the integrity of fine cellular structures and for subsequent quantitative analyses in Imaris software. Imaging is performed with a 40× PlanApo oil-immersion objective (numerical aperture 1.4), achieving a resolution of 1,024 × 1,024 pixels. The pixel dimensions are 0.156 μm × 0.156 μm, with a voxel depth of 0.8 μm. Two channels are acquired during imaging: 405 nm for DAPI and 647 nm for Alexa Fluor 647 antibody. Optimize the gain, offset, and laser power to maximize signal-to-noise ratio while minimizing photobleaching. Under our experimental conditions for IBA1 acquisition, the channel detector gain was set to 720, the offset to -40, and the laser power to 6.5. However, it is important to note that these settings are specific to this experiment and do not represent a standardized protocol.
The Z-stack function in LSM works as follows:
Set First/Last Mode: Define the upper and lower limits to specify the Z-axis range used in 3D imaging.
Note: Select the channel of interest and click “live” mode to preview the sample; then, set the start and end of the Z-stack with this mode.
Z-stack capture: Once the limits have been settled, press “start experiment” to acquire the Z-stack.
D. 3D reconstruction and image pre-processing
The protocol outlined below is tailored for image analysis using the commercial Imaris software. For those new to the software, please refer to online video tutorials and webinars available at http://www.bitplane.com/learning and https://imaris.oxinst.com/homeschool.
1. Upload the image and display adjustment (Video 1)
a. Upload Z-stack serial images of the microglia in Imaris 9.6. The software will automatically generate 3D images.
Note: Imaris can convert single images or a series from .lsm file format to the required .ims format
b. Select the channel corresponding to the signal and adjust the background fluorescence using Channel Max Intensity to minimize background noise and ensure accurate quantification.
Video 1. Automatic three-dimensional reconstruction of microglial cells using Imaris software
2. Imaris Filament Tracer (Figures 1–3 and Video 2)
a. Add a new filament by clicking on the leaf icon.
b. Click on the blue arrow to proceed to the next page and select the channel of interest.
c. Manually adjust the parameter “Largest Diameter” in the starting point section.
Note: In the slice view (two-dimensional viewer), measure several soma distances to adjust the largest diameter accordingly.
d. Correct any inaccuracies in the dendrite beginning points manually by placing the point at the center of the soma or removing it as needed.
e. Click on the green double arrow to finish the rendering.
f. Select the Edit tab and remove any filaments not suitable for further analysis using the Delete function
Note: Remove all filaments located on the tissue edge of the Z-stack. For falsely fused filaments, separate them by selecting a specific point and using the delete function. If separation is not feasible, remove those filaments from the analysis.
g. Click on the Results tab and select the specific values you want to analyze. Export the data for further analysis.
Note: For the comparison of filaments based on treatment and genotype, dendrite branch level and dendrite volume have proven to be efficient and sensitive parameters.
Figure 1. Graphical representation of dendrite branch level and dendrite volume. A. The numbers represent each ramification level, with each dendrite branch level distinguished by different colors for clear differentiation. B. The blue-shaded area (starting from the darker blue circle) highlights the dendrite volume, illustrating its distribution along the dendritic structure.
Figure 2. Three-dimensional reconstruction of microglial filaments. A. Select the leaf icon to initiate filament rendering. B. Click on the blue arrow to proceed to parameter adjustments. C. Choose the source channel and manually adjust the “largest diameter” value. D. De-select spine detection, then click the green double arrow to complete the reconstruction. E. Switch to the Pencil tab for editing filaments. F. Navigate to the Results tab and select the parameters of interest from the drop-down menu for further analysis.
Figure 3. Three-dimensional editing of microglial filaments. Microglial cells located away from the tissue edges were selected for further editing. A. Filaments automatically reconstructed by Imaris software. B. Filaments post-editing, using surface rendering as a guide for accurate refinement.
Video 2. Three-dimensional reconstruction and manual editing of microglial filaments using the Imaris Filament Tracer.
3. Soma volume (Figures 4–5)
a. To create a surface, click on the blue gem icon.
b. In the panel below, choose “skip automatic creation, edit manually.”
c. Navigate through the Z-stacks to locate the soma of interest. Use the draw tool to trace the outline of the soma on at least three different levels.
Note: Ensure you select Z-stack slices that provide the clearest image of the soma.
d. Once done, click on create surface, and the software will generate and analyze the selected surfaces.
e. Navigate to the Results section, select the relevant parameters, and choose the measurement you require, such as soma volume.
Figure 4. Three-dimensional reconstruction of soma volume of microglial cells. A. Select the surface rendering tool. B. Skip automatic creation and proceed with manual editing. C. Employ the draw tool to outline the microglial somas, then create the corresponding surfaces for volume analysis.
Figure 5. Three-dimensional reconstruction of microglial soma volume. Z-stack images of microglial cells labeled with the IBA1 antibody. A. Surface and filament rendering displaying the microglial cells selected for analysis. B. Magnified view of the region in panel A. C. Manual contouring of microglial somas (dotted blue lines) across individual slices of the Z-stack D. Merged soma contours from all Z-stack slices for the selected microglial cells. E. 3D reconstruction of microglial somas from the merged contours. F. Each reconstructed soma corresponds to a filament reconstruction, previously selected for analyses. Scale bars = 20 μm in A, D, E, F; 10 μm in B, C.
4. Imaris 3D surface (Figure 6 and Video 3)
a. Add a new Surface by clicking on the blue gem icon.
b. Click on the blue arrow to proceed, then select the channel of interest.
c. Adjust the Surfaces Detail parameter to increase or decrease surface precision.
Note: If the Smooth option is selected, surface detail will be calculated automatically. Manually increasing this value will reduce surface detail.
d. Manually adjust the Threshold (Absolute Intensity) parameter until the signal of interest is fully captured.
e. Modify the Filter Seed Point parameter to ensure all desired filaments are included.
f. Click Finish to complete the filament rendering.
Figure 6. Three-dimensional reconstruction of microglial surfaces. A. Select the source channel and manually adjust the surface detail parameter. B. Adjust the threshold to ensure that all signals of interest are captured. C. Adjust the filter seed points to include the desired filaments in the final reconstruction.
Video 3. Three-dimensional surface reconstruction of microglial cells using Imaris software.
Data analysis
Custom Python scripts were developed to facilitate the analysis of Imaris outputs. First, export the relevant .csv files containing the desired parameters from Imaris for further processing (Figure 7).
Note: Imaris generates two .csv files for each Z-stack, one containing dendrite volume data and the other containing branch-level data for the analyzed cells.
For each Z-stack .csv file, run the DendriteVolume.py and DendriteBranchLevel.py scripts as follows:
-To select specific output files for comparison:
python3 DendriteScript.py pathtocsvfile1 pathtocsvfile2 pathtocsvfile3 …
-To analyze all samples within a folder:
python3 DendriteScript.py pathtofolder/*
-If no input files are specified, the script will attempt to analyze all .csv files located in the same directory as the script.
The script will output the total dendrite volume (Dendrite Volume) and the total number of branches for each branch level (Dendrite Branch Level) for the same microglial cell (Filament ID).
1. Dendrite volume output: The first line displays the name of the .csv file (e.g., 2248a_zstack_40x_zoom1_Detailed.csv). Below this, two columns are presented: the first lists the microglial cell IDs, and the second shows the total dendrite volume.
2. Dendrite branch level output: Similarly, the first line shows the name of the .csv file (e.g., 2284a_zstack_40x_zoom1_Detailed.csv). The second line represents the Filament ID, followed by two columns: one for the branch level (with 1 representing the first branch from the soma) and another for the number of branches at each level.
Figure 7. Output examples of dendrite volume and dendrite branch Level. The identifier 100000034 corresponds to one of the three microglial cells analyzed in the 2248a z-stack. The orange square indicates the total measured dendrite volume of the cell, while the green square represents the number of branches at each branch level (e.g., 5 branches for the first level, 23 branches for the second level).
Validation of protocol
This protocol has been used and validated in Figure 3f–j of the following research article:
Puighermanal, E. et al. [13]. Cannabidiol ameliorates mitochondrial disease via PPARγ activation in preclinical models. Nat Commun.
In this research article, we used the protocol described here to analyze 3D microglial morphology (Figure 8A) in the external globus pallidus, a highly affected brain region in Leigh syndrome mouse models. We demonstrated that mice with conditional Ndusf4 ablation in Gad2-espressing neurons exhibited morphometric remodeling of microglia resembling the activated phenotype. Specifically, microglial cells in mutant mice displayed a larger soma size (Figure 8B) and dendrite volume (Figure 8C) compared to control mice, along with fewer branch levels and fewer branches per level (Figure 8D). Total filament length was not altered (Figure 8E). Data represented in Figure 8 are extracted from Puighermanal et al. [13].
Figure 8. Morphometric analyses of microglia in a Leigh syndrome mouse model that courses with microgliosis. A. Imaris 3D surface reconstruction of a microglial cell. B. Quantification of soma volume/cell of IBA1+ cells (n = 9 control and 7 Gad2:Ndufs4cKO mice, two-sided t-test, ****p < 0.0001). C. Quantification of total dendrite volume/cell of IBA1+ cells (n = 9 control and 7 Gad2:Ndufs4cKO mice, two-sided t-test, ****p < 0.0001). D. Distribution plot of microglial ramification (n = 9 control and 7 Gad2:Ndufs4cKO mice, two-way ANOVA repeated measures, ****p < 0.0001). E. Quantification of total dendrite length/cell of IBA1+ cells (n = 9 control and 7 Gad2:Ndufs4cKO mice, two-sided t-test). All data are presented as mean values ± SEM. Veh vehicle, CT control mice, cKO Gad2:Ndufs4cKO mice. Scale bar: 8 μm.
General notes and troubleshooting
Troubleshooting
Problem 1: Densely packed cells make it difficult to reconstruct individual microglial filaments in 3D.
Possible cause: High cell density in the region of interest, especially in central areas of the sample and/or due to a neuroinflammatory environment, which increases microglial proliferation and clustering.
Solutions:
1. Select regions with more isolated cells for imaging. Focus on peripheral areas of the sample where cell density is typically lower.
2. Manually exclude overlapping or crowded cells during analysis by erasing them in Imaris. This approach allows for a cleaner reconstruction of the remaining cells.
3. In samples with neuroinflammation, consider imaging regions with lower microglial density to reduce complexity in filament reconstruction.
Problem 2: Differences in intensity and morphology of microglial cells between experimental groups impact imaging consistency.
Possible cause: Confocal settings (e.g., laser power, gain, offset) optimized for one group may not work well for others due to condition-dependent variability.
Solutions:
1. Test and optimize imaging parameters (e.g., laser intensity, detector gain) across all experimental groups before starting batch imaging.
2. Adjust settings to achieve a balance that provides consistent signal-to-noise ratios and preserves cell morphology across conditions.
Acknowledgments
The authors apologize to any colleagues whose work was not discussed in this paper. The authors greatly thank the Microscopy Imaging Platform at Autonomous University of Barcelona for continuous technical support with the Imaris software and Kevin Aguilar for technical advice.
This work was supported by the MINECO Ramon y Cajal (RYC2020-029596-I) and MICINN (PID2021-125079OA-I00) grants awarded to E.P., as well as the 2024 FI-100645 fellowship granted to G.vdW.
Author Contributions
E.P. and M.dD conceptualized and planned the study. M.dD performed immunofluorescence studies and image analysis. A.dD. and M.dD. created the data analysis pipeline and analyzed the pertaining data, A.K. generated, rendered and edited the final audiovisual content reported. M.dD., A.K., and E.P. wrote the manuscript. G.vdW. edited the manuscript.
Competing interests
The authors declare no conflict of interest.
Ethical considerations
All animal experiments were performed with the approval of the ethical committee at Autonomous University of Barcelona (CEEAH) and Generalitat de Catalunya (DMAH).
References
Aguilar, K., Comes, G., Canal, C., Quintana, A., Sanz, E. and Hidalgo, J. (2022). Microglial response promotes neurodegeneration in the Ndufs4 KO mouse model of Leigh syndrome. Glia. 70(11): 2032–2044. https://doi.org/10.1002/glia.24234
Lee, J. W., Chun, W., Lee, H. J., Kim, S. M., Min, J. H., Kim, D. Y., Kim, M. O., Ryu, H. W. and Lee, S. U. (2021). The Role of Microglia in the Development of Neurodegenerative Diseases. Biomedicines. 9(10): 1449. https://doi.org/10.3390/biomedicines9101449
Leyh, J., Paeschke, S., Mages, B., Michalski, D., Nowicki, M., Bechmann, I. and Winter, K. (2021). Classification of Microglial Morphological Phenotypes Using Machine Learning. Front Cell Neurosci. 15: e701673. https://doi.org/10.3389/fncel.2021.701673
Doorn, K. J., Goudriaan, A., Blits‐Huizinga, C., Bol, J. G., Rozemuller, A. J., Hoogland, P. V., Lucassen, P. J., Drukarch, B., van de Berg, W. D., van Dam, A., et al. (2013). Increased Amoeboid Microglial Density in the Olfactory Bulb of Parkinson's and Alzheimer's Patients. Brain Pathol. 24(2): 152–165. https://doi.org/10.1111/bpa.12088
Green, T. R. F., Murphy, S. M. and Rowe, R. K. (2022). Comparisons of quantitative approaches for assessing microglial morphology reveal inconsistencies, ecological fallacy, and a need for standardization. Sci Rep. 12(1): e1038/s41598–022–23091–2. https://doi.org/10.1038/s41598-022-23091-2
Clarke, D., Crombag, H. S. and Hall, C. N. (2021). An open-source pipeline for analysing changes in microglial morphology. Open Biol. 11(8): e210045. https://doi.org/10.1098/rsob.210045
Heindl, S., Gesierich, B., Benakis, C., Llovera, G., Duering, M. and Liesz, A. (2018). Automated Morphological Analysis of Microglia After Stroke. Front Cell Neurosci. 12: e00106. https://doi.org/10.3389/fncel.2018.00106
Verdonk, F., Roux, P., Flamant, P., Fiette, L., Bozza, F. A., Simard, S., Lemaire, M., Plaud, B., Shorte, S. L., Sharshar, T., et al. (2016). Phenotypic clustering: a novel method for microglial morphology analysis. J Neuroinflammation. 13(1): 153. https://doi.org/10.1186/s12974-016-0614-7
York, E. M., LeDue, J. M., Bernier, L. P. and MacVicar, B. A. (2018). 3DMorph Automatic Analysis of Microglial Morphology in Three Dimensions fromEx VivoandIn VivoImaging. eNeuro. 5(6): ENEURO.0266–18.2018. https://doi.org/10.1523/eneuro.0266-18.2018
Gavrin, A. and Fedorova, E. (2015). Quantification of the Volume and Surface Area of Symbiosomes and Vacuoles of Infected Cells in Root Nodules of Medicago truncatula. Bio Protoc. 5(23): e1665. https://doi.org/10.21769/bioprotoc.1665
Ohnishi, Y. and Okamoto, T. (2015). Microscopic Observation, Three-dimensional Reconstruction, and Volume Measurements of Sperm Nuclei. Bio Protoc. 5(7): e1437. https://doi.org/10.21769/bioprotoc.1437
Bolea, I., Gella, A., Sanz, E., Prada-Dacasa, P., Menardy, F., Bard, A. M., Machuca-Márquez, P., Eraso-Pichot, A., Mòdol-Caballero, G., Navarro, X., et al. (2019). Defined neuronal populations drive fatal phenotype in a mouse model of Leigh syndrome. eLife. 8: e47163. https://doi.org/10.7554/elife.47163
Puighermanal, E., Luna-Sánchez, M., Gella, A., van der Walt, G., Urpi, A., Royo, M., Tena-Morraja, P., Appiah, I., de Donato, M. H., Menardy, F., et al. (2024). Cannabidiol ameliorates mitochondrial disease via PPARγ activation in preclinical models. Nat Commun. 15(1): 7730. https://doi.org/10.1038/s41467-024-51884-8
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Article Information
Publication history
Received: Oct 16, 2024
Accepted: Jan 8, 2025
Available online: Jan 26, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Category
Neuroscience > Nervous system disorders
Neuroscience > Cellular mechanisms > Microglia
Biological Sciences > Biological techniques
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5,219 | https://bio-protocol.org/en/bpdetail?id=5219&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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This is an In Press version of the protocol that has not yet been assigned to an issue.
Peer-reviewed
Quantification of Neuromuscular Junctions in Zebrafish Cranial Muscles
RG Ritika Ghosal
JE Johann K. Eberhart
In Press, Available online: Jan 26, 2025
DOI: 10.21769/BioProtoc.5219 Views: 29
Reviewed by: Ivonne SehringAmr Galal Abdelraheem Ibrahim Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Frontiers in Physiology Feb 2023
Abstract
Communication between motor neurons and muscles is established by specialized synaptic connections known as neuromuscular junctions (NMJs). Altered morphology or numbers of NMJs in the developing muscles can indicate a disease phenotype. The distribution and count of NMJs have been studied in the context of several developmental disorders in different model organisms, including zebrafish. While most of these studies involved manual counting of NMJs, a few of them employed image analysis software for automated quantification. However, these studies were primarily restricted to the trunk musculature of zebrafish. These trunk muscles have a simple and reiterated anatomy, but the cranial musculoskeletal system is much more complex. Here, we describe a stepwise protocol for the visualization and quantification of NMJs in the ventral cranial muscles of zebrafish larvae. We have used a combination of existing ImageJ plugins to develop this methodology, aiming for reproducibility and precision. The protocol allows us to analyze a specific set of cranial muscles by choosing an area of interest. Using background subtraction, pixel intensity thresholding, and watershed algorithm, the images are segmented. The binary images are then used for NMJ quantification using the Analyze Particles tool. This protocol is cost-effective because, unlike other licensed image analyzers, ImageJ is open-source and available free of cost.
Key features
• Immunostaining neuromuscular junctions in alcohol-exposed zebrafish.
• Quantification of presynaptic and postsynaptic terminals in cranial muscles of zebrafish larvae.
• Analysis of size and distribution of NMJs in cranial muscles of zebrafish larvae.
Keywords: Neuromuscular junctions Zebrafish Cranial nerves FIJI/ImageJ Ethanol Automated particle quantification
Graphical overview
Background
In vertebrates, the muscles of the face and neck are innervated by cranial motor neurons [1]. Communication between the brain and muscle is mediated through a specialized synapse known as the neuromuscular junction (NMJ). NMJs typically comprise three components: 1) the axonal end of the motor neuron, forming the presynaptic terminal; 2) the muscle endplate, forming the postsynaptic terminal; and 3) a synaptic cleft that lies between these two terminals. Neuromuscular communication occurs when acetylcholine, released from the presynaptic terminal into the synaptic cleft, binds acetylcholine receptors clustered in the postsynaptic terminal [2]. Alteration in NMJ morphology or distribution can lead to reduced synaptic transmission and result in neuromuscular disease. Zebrafish has emerged as an excellent model to study neuromuscular pathologies [3]. However, characterization and quantification of NMJs in the cranial muscles of zebrafish can be particularly challenging due to the anatomical complexity of the head.
Previous studies involving the quantification of NMJs in the cranial muscles were primarily carried out manually. Manual counting is arduous and can lead to individual bias and subjectivity. Some recent studies have carried out automated quantification using image analysis tools. However, most of these studies were carried out in somites of the zebrafish trunk region [4,5]. Unlike the head, the muscles of the trunk have a simpler, reiterated anatomy. Another study employed Imaris software for surface rendering with automatic thresholding to characterize the NMJs in the cranial musculature [6]. This method is useful for studying the number, shape, and architecture of NMJs. However, this also requires buying licensed software, making it inaccessible to some labs. Also, most of these studies use auto-thresholding, which can change the cutoff value for thresholding in individual images, unintentionally introducing errors [7].
Here, we developed a protocol for NMJ visualization and quantification in the cranial muscles using FIJI/ImageJ software, which is free of cost [8]. We immunolabel the presynaptic and postsynaptic terminals using SV2 antibody and alpha-bungarotoxin, respectively. We then capture high-resolution images of NMJs using confocal microscopy. Using the Area Selection tool, we crop out a region of specific dimensions containing the muscles of interest. We then use the Rolling Ball Background Subtraction plugin, Global Thresholding, and Watershed plugin to segment the images. Instead of auto-thresholding, we set the same cutoff value for thresholding each image, ensuring uniformity across all images. The number of particles in the segmented images is then counted using the Analyze Particles tool. We have used this protocol to investigate the effect of alcohol on NMJ count in a specific set of cranial muscles. However, the method can be applied to other cranial and trunk muscles of zebrafish. The method can also be used to quantify NMJs in other organisms like chickens and mice. Additionally, we used this method with 20× magnification for NMJ counting, but the method could readily be adapted for analyzing images of other magnifications. This method is broadly applicable to quantify other fluorescently labeled particles such as apoptotic cells, proliferating cells, or cells in general by counting DAPI-stained nuclei.
Materials and reagents
Biological materials
1. Zebrafish (Animal husbandry, Eberhart Lab, University of Texas at Austin, USA)
Reagents
1. Alpha-bungarotoxin (α-BTX) Alexa Fluor 647 conjugated (Invitrogen, catalog number: B35450)
2. Bovine serum albumin (BSA) (Sigma-Aldrich, catalog number: A4503, CAS: 9048-46-8)
3. Calcium chloride dihydrate (CaCl2·2H2O) (Sigma, catalog number: C8106, CAS: 10035-04-8)
4. Dimethyl sulfoxide (DMSO) (EMD Millipore Corp., catalog number: 317275, CAS: 67-68-5)
5. Ethanol (PHARMO, Ethyl Alcohol 100%, catalog number: 111000200, CAS: 64-17-5)
6. Goat anti-mouse IgG, Alexa Fluor 488 (secondary antibody) (Invitrogen, catalog number: A-11029)
7. Magnesium sulfate heptahydrate (MgSO4·7H2O) (Sigma-Aldrich, catalog number: 230391, CAS: 10034-99-8)
8. MESAB/Tricaine methanesulfonate (Syndel, SYNCAINE/MS 222, FDA approved)
9. Methanol (Fisher Chemical, catalog number: A412-4, CAS: 67-56-1)
10. Methylcellulose (Sigma-Aldrich, catalog number: M0387, CAS: 9004-67-5)
11. Normal goat serum (NGS) (Jackson ImmunoResearch Laboratories, catalog number: 005-000-121)
12. Paraformaldehyde (PFA) (Alfa Aesar, catalog number: A11313, CAS: 30525-89-4)
13. Phosphate-buffered saline (PBS) (Sigma-Aldrich, catalog number: P5368-10PAK)
14. Potassium chloride (KCl) (Fisher chemicals, catalog number: P217, CAS: 7447-40-7)
15. Potassium phosphate dibasic (K2HPO4) (Sigma-Aldrich, catalog number: P3786, CAS: 7758-11-4)
16. Sodium bicarbonate (NaHCO3) (Sigma-Aldrich, catalog number: S5761, CAS: 144-55-8)
17. Sodium chloride (NaCl) (CHEM-IMPEX INT’L INC., catalog number: 30070, CAS: 7647-14-5)
18. Sodium phosphate dibasic (Na2HPO4) (Acros organics, catalog number: 42437, CAS: 7558-79-4)
19. Synaptic vesicle 2A (SV2) antibody (Developmental Studies Hybridoma Bank, Antibody Registry ID: AB_2315387)
20. Triton X-100 (Triton X) (MP Biomedicals, catalog number: 194854, CAS: 9002-93-1)
Solutions
1. 10× PBS (See Recipes)
2. 1× PBT (See Recipes)
3. 4% PFA (See Recipes)
4. 25% methanol (See Recipes)
5. 50% methanol (See Recipes)
6. 75% methanol (See Recipes)
7. 20× embryo medium stock (See Recipes)
8. 500× sodium bicarbonate (See Recipes)
9. Embryo medium (See Recipes)
10. 1% ethanol (See Recipes)
11. Incubation buffer (See Recipes)
12. Blocking buffer (See Recipes)
13. Ethyl 3 Aminobenzoate methyl sulfonate salt (MESAB)/tricaine solution (See Recipes)
Recipes
1. 10× PBS
Reagent Final concentration Quantity or Volume
Phosphate-buffered saline 10× 1 packet
ddH2O n/a Up to 100 mL
Total n/a 100 mL
2. 1× PBT
Reagent Final concentration Quantity or Volume
10× PBS 1× 5 mL
Triton X 0.5% (v/v) 250 µL
ddH2O n/a Up to 50 mL
Total n/a 50 mL
3. 4% PFA
*Note: Add PFA to 400 mL of ddH2O and heat and stir in a fume hood until the solution clears (do not heat at more than 60 °C). Add 50 mL of 10× PBS and adjust the pH to 7.4. Add ddH2O up to a total of 500 mL.
Reagent Final concentration Quantity or Volume
10× PBS 1× 50 mL
PFA 4% (w/v) 20 g
ddH2O n/a 450 mL*
Total n/a 500 mL
Store 4% PFA at -20 °C.
4. 25% methanol
Reagent Final concentration Quantity or Volume
Methanol 25% (v/v) 1 mL
1× PBT 75% (v/v) 3 mL
Total n/a 4 mL
5. 50% methanol
Reagent Final concentration Quantity or Volume
Methanol 50% (v/v) 2 mL
1× PBT 50% (v/v) 2 mL
Total n/a 4 mL
6. 75% Methanol
Reagent Final concentration Quantity or Volume
Methanol 75% (v/v) 3 mL
1× PBT 25% (v/v) 1 mL
Total n/a 4 mL
7. 20× embryo medium stock
Reagent Final concentration Quantity or Volume
NaCl 0.3 M 17.5 g
KCl 10.06 mM 0.75 g
CaCl2·2H2O 19.72 mM 2.9 g
K2HPO4 2.35 mM 0.41 g
Na2HPO4 1 mM 0.142 g
MgSO4·7H2O 19.88 mM 4.9 g
ddH2O n/a Up to 1 L
Total n/a 1 L
Filter-sterilize the solution.
8. 500× sodium bicarbonate
Reagent Final concentration Quantity or Volume
NaHCO3 0.35 M 0.3 g
ddH2O n/a 10 mL
9. Embryo medium
Reagent Final concentration Quantity or Volume
20× embryo medium stock 1× 50 mL
500× sodium bicarbonate 1× 2 mL
ddH2O n/a 948 mL
Total n/a 1 L
10. 1% Ethanol
Reagent Final concentration Quantity or Volume
Ethanol 1% (v/v) 400 µL
Embryo medium 99% (v/v) 39.6 mL
Total n/a 40 mL
11. Incubation buffer
Reagent Final concentration Quantity or Volume
10× PBS 1× 5 mL
BSA 1% (w/v) 500 mg
DMSO 1% (v/v) 500 µL
Triton X-100 0.5% (v/v) 250 µL
ddH2O n/a Up to 50 mL
Total n/a 50 mL
12. Blocking buffer
Reagent Final concentration Quantity or Volume
Incubation buffer 96% (v/v) 960 µL
Normal goat serum 4% (v/v) 40 µL
Total n/a 1,000 µL
13. MESAB/tricaine solution
Reagent Final concentration Quantity or Volume
Na2HPO4 0.8 % (w/v) 4 g
MESAB salt 0.4 % (w/v) 2 g
ddH2O n/a Up to 500 mL
Total n/a 500 mL
Adjust pH between 7.0 and 7.2.
Laboratory supplies
1. Petri dishes 100 mm × 20 mm (Falcon, catalog number: 353003)
2. Micropipettes (Sigma-Aldrich, catalog number: FA10006M-1EA)
3. Pipette pumps (Fisher Scientific, catalog number: 13-683C)
4. 1.7 mL microcentrifuge tubes (Fisher Scientific, catalog number: 07-200-535)
Equipment
1. Confocal microscope (Zeiss, model: LSM 980)
2. Stereoscope (Leica, model: KL 300 LED)
Software and datasets
1. ImageJ2 v2.14.0/1.54f (07/07/2023, available free of cost from https://fiji.sc/)
2. Prism v9.5.0 (GraphPad, 12/06/2022)
Procedure
Here, we describe a stepwise protocol to visualize and quantify the NMJs in the cranial muscles of zebrafish larvae at 4 days post fertilization. This protocol has been used to compare the number of NMJs in ethanol-exposed and unexposed zebrafish [9]. The protocol can be adapted for comparing NMJs at other times, in other conditions, and in other muscle(s).
A. Sample preparation
1. Alcohol treatment of zebrafish embryos
a. Harvest wildtype zebrafish embryos and incubate at 28 °C in 40 mL of embryo medium (see Recipes) in a 100-mm Petri dish with a maximum number of 100 embryos per dish. Allow the embryos to develop until shield stage, which is at about 6 h post fertilization (6 hpf). Zebrafish staging has been previously described [10].
b. At 6 h post-fertilization, freshly prepare 40 mL of 1% ethanol in embryo medium (see Recipes). Replace the embryo medium in the experimental dish with the freshly prepared 1% ethanol solution. Leave the control dish with embryo medium untreated. Place the experimental and control dishes in the incubator at 28 °C and allow the embryos to develop until 4 days post fertilization (4 dpf). Keep the dishes with the developing zebrafish embryos clean by periodically (daily at a minimum) removing any dead embryos or chorions shed during hatching.
2. Fixation
a. At 4 dpf, harvest 15 fish each from the control and experimental dishes. To harvest zebrafish, euthanize using MESAB/tricaine (see Recipes) in embryo medium. Using a graduated dropper, add MESAB/tricaine to a final concentration of 200–300 mg/L. Transfer the euthanized zebrafish from the control and experimental dishes to 1.7 mL microcentrifuge tubes using a pipette pump.
b. Remove any excess liquid from the microcentrifuge tubes using a pipette pump.
c. Add 1 mL of 4% PFA solution (see Recipes) and place the microcentrifuge tubes on a nutator at 4 °C for overnight fixation.
Caution: PFA is toxic and should be disposed of according to institutional regulations. Proper care must be taken while working with PFA (e.g., use of gloves and working in a chemical hood).
d. After overnight fixation, remove the 4% PFA solution from the tubes and wash the zebrafish larvae with 1 mL of 1× PBS for 5 min on the nutator at room temperature (wash at least twice to ensure complete removal of PFA).
Note: After the wash step, the zebrafish are ready for the immunohistochemical staining. However, to store the zebrafish larvae until later use, carry out methanol dehydration as described in the next step.
3. Methanol dehydration for long-term storage (optional)
a. For methanol dehydration, prepare 25%, 50%, and 75% methanol solutions in 1× PBT (see Recipes). Dehydrate larvae with serially diluted methanol solutions (1× PBT, 25%, 50%, 75%, and then 100% methanol) by washing the larvae in 1 mL of each solution for 5 min on the nutator at room temperature. Replace 100% methanol with 1 mL of cold 100% methanol. Store the zebrafish larvae at -20 °C.
Pause point: The dehydrated larvae can be stored in 70% methanol solution at 4 °C for a week or in 100% methanol at -20 °C for a maximum of 6 months.
b. Rehydrate the zebrafish larvae stored in methanol at -20 °C. Wash once for 5 min with the serially diluted methanol solutions (75%, 50%, 25%, and then 1× PBT) on the nutator at room temperature.
B. Immunohistochemistry for NMJs in cranial muscles
1. Freshly prepare incubation buffer (see Recipes) and wash four times in 1 mL of incubation buffer for 20 min each on the nutator at room temperature.
2. Freshly prepare blocking buffer (see Recipes). Wash the zebrafish once with 500 µL of blocking buffer for 30 min on the nutator at room temperature.
3. Prepare 250 µL of primary antibody solution in blocking buffer. For labeling the presynaptic terminals, use primary antibody against synaptic vesicle 2A (see Reagents) to get a final concentration of 5 µg/mL. Replace the blocking buffer with primary antibody solution and place the microcentrifuge tubes on a nutator with gentle rocking at 4 °C for overnight incubation.
4. The following day, remove the primary antibody solution and wash the zebrafish larvae four times with 1 mL of incubation buffer for 20 min each on the nutator at room temperature.
5. Wash once with 500 µL of blocking buffer for 30 min on the nutator at room temperature.
6. Prepare 250 µL of anti-mouse IgG Alexa Fluor 488 secondary antibody solution in blocking buffer at 1:500 dilution by adding 0.5 µL of secondary antibody to 250 µL of blocking buffer. Also, add 1 µL of alpha-bungarotoxin (α-BTX) Alexa Fluor 647 conjugated primary antibody to obtain a 1:250 dilution (see Reagents for details on antibodies).
7. Replace the blocking buffer with the antibody solution and cover the tubes with aluminum foil to protect from the light. Place the tubes on an incubator at 4 °C for overnight incubation with gentle rocking.
8. The following day, remove the antibody solution from the tubes and wash three times with 1 mL of 1× PBT for 5 min each on the nutator at room temperature.
9. Fix the immunostained zebrafish larvae for 20 min by washing in 4% PFA on the nutator at room temperature.
10. Wash twice in 1 mL of 1× PBT for 5 min each on the nutator at room temperature.
11. Replace the 1× PBT solution with 1 mL of 1× PBS solution. The zebrafish larvae are ready to be imaged.
Pause point: Immunostained larvae in 1× PBS can be stored at 4 °C covered in foil (protected from light) for a week before imaging.
Note: It is recommended that they are imaged as soon as possible after immunostaining to prevent loss of fluorescence.
C. Confocal imaging
Mount the immunolabeled zebrafish to collect ventral images in 0.2% agarose and 3% methylcellulose as previously described [11]. To mount the samples, add a drop of methylcellulose using a wooden applicator in the middle of a triple-bridged coverslip. Using a pipette pump, transfer the immunolabeled zebrafish from 1× PBS to 0.2% agarose. Next, transfer the zebrafish along with some agarose solution to the methylcellulose drop. For an upright microscope, orient the zebrafish in a ventral-upward position using a capillary poker and carefully place the coverslip on the top. Acquire bidirectional confocal Z-stacks at 20× magnification. We used the Zeiss LSM 980 microscope with 20× objective lens, 1,024 × 1,024 frame size, and 4× averaging. Confocal stacks were Z-projected with maximum intensity as shown in Figure 1 for further image analyses described in section D.
Note 1: Make sure to image through the entire depth of the brachial region to capture all muscle fibers. Most ventral cranial muscles, such as the intermandibularis posterior and interhyoideus and hyohyoideus inferior muscles, are fairly superficial. However, the intermandibularis anterior, hyohyoideus superior, sternohyoideus, and adductor mandibularis muscles are much deeper. Make sure to image through their depth to capture the entire muscle. For 4 dpf zebrafish, a depth of 60 µm should be sufficient to capture all ventral branchial muscles shown in Figure 1A.
Note 2: Make sure that all images for analysis are captured using the same confocal settings such as laser intensity, gain, frame size, averaging, etc.
Note 3: Also, the distance between two consecutive Z-slices should be the same across all images for comparison and should not be more than 2 µm for looking at the NMJs.
Figure 1. Confocal images of ventral cranial muscles immunostained with synaptic vesicle 2A (SV2) and alpha-bungarotoxin (α-BTX) in zebrafish at 4 dpf. A, D. Merged channels with SV2-stained presynaptic terminals (in green) and α-BTX-stained postsynaptic terminals (in magenta) in untreated and ethanol-treated zebrafish. Ventral cranial muscles in zebrafish larvae at 4 dpf, as previously described [12], are shown in A: M.IMA: intermandibularis anterior muscle; M.IMP: intermandibularis posterior muscle; M.IH: interhyoideus muscle; M.AM: adductor mandibulae muscle; M.HHI: hyohyoideus inferior muscle; M.HHS: hyohyoideus superior muscle; M.SH: sternohyoideus muscle. B, E. SV2-labeled presynaptic terminals in untreated and alcohol-exposed zebrafish. C, F. α-BTX-labeled postsynaptic terminals in untreated and alcohol-exposed zebrafish. Scale bar = 100 µm.
D. Image analysis using FIJI/ImageJ software
To study the size and distribution of presynaptic and postsynaptic terminals in the ventral cranial muscles of zebrafish larvae, FIJI/ImageJ software can be used. Analyses can be carried out for one or more cranial muscles of interest. In this example, the quantification of the NMJs in four cranial muscles, the intermandibularis anterior, intermandibularis posterior, interhyoideus, and hyohyoideus inferior muscles has been carried out.
1. Area selection to pick the muscles of interest:
a. Open the image file with ImageJ software. To Z-project the image, select the image and choose Image → Stacks → Z project from the ImageJ menu options. Select Max Intensity as the Projection type and click OK on the ZProjection dialog box.
b. To pick the muscles of interest, use the rectangular selection tool to crop out a region of 300 × 180 µm^2 (= 750 × 450 in pixel units) containing intermandibularis anterior, intermandibularis posterior, interhyoideus, and hyohyoideus inferior muscles. To specify the dimensions for the area of interest, choose Edit → Selection → Specify. The Specify dialog box will appear. Check the Scaled unit (microns) box and enter 300 and 180 as the Width and Height, respectively, as illustrated in Figure 2A. Other sizes can be used depending on the researcher’s specific interests. Click OK to proceed.
c. Using the arrow tool, drag the rectangle of the specified dimensions to the desired location to select the muscles of interest. For instance, here, the rectangle outlines the intermandibularis anterior, intermandibularis posterior, interhyoideus, and hyohyoideus inferior muscles (Figure 2A).
d. To crop out the selected region, choose Image → Crop or Image → Duplicate. Only the selected region will be included in the duplicated image (Figure 2B). All further steps will be carried out on this duplicated or cropped image with the muscles of our interest.
Figure 2. Area selection to select a region of specific dimensions. A. Z-projected image with the rectangular selection (in yellow) of 300 µm in width and 180 µm in height. The dialog box on the right shows the values specified for width and height in scaled units (microns). B. Cropped region of the image with 300 × 180 µm^2 selected area for analysis. Scale bar = 100 µm.
2. Background subtraction: To correct the uneven background illumination, use the Rolling ball background subtraction plugin.
a. To apply the method, select Process → Subtract Background from the ImageJ menu options. The Subtract Background dialog box will appear. Enter the Rolling ball radius and press OK.
b. To determine the Rolling ball radius, measure the length of the largest object (i.e., NMJ) by using the Line selection tool on ImageJ. Draw a line connecting two dots that defines the length of the largest object on the image and select Analyze → Measure to get the length in microns for a scaled image (Figure 3). Here, the largest length was found to be ~10 (9.199) µm.
c. For this image, pixel size = 0.4 µm in length and breadth. Thus, 10 µm = 25 pixels. Enter 25 as the Rolling ball radius in pixels and click OK. A Process Stack? dialog box will appear. To apply the background subtraction to both SV2 and α-BTX channels, select Yes to Process all images? on the Process Stack? dialog box. Figure 4 shows the images before and after correcting their background illumination using the Rolling ball background subtraction plugin.
Figure 3. Determining the rolling ball radius by measuring the length of the largest object on the image. A. The red arrow shows the line selection tool. B. Line drawn with the line selection tool to denote the length of the largest object on the image. C. Result table displaying the length of the line drawn as 9.199 µm (highlighted by the red rectangle).
Note 1: To find the pixel size of your image, select the image and choose Image → Properties from the ImageJ menu options.
Note 2: In theory, the rolling ball radius is determined by measuring the length of the largest object or particle in the image that is considered a true signal and not part of the background. However, in practice, the optimum radius can be worked out by trying different values using the Preview option on the Subtract Background window. If the value is too large, it might not fix the uneven background; on the other hand, a very small radius can take away parts of the real signal (synaptic terminals in this case).
Note 3: The rolling ball radius can be different for different channels. So, determine the radius for both channels separately using step D2b. However, here, we found that the largest particles on both channels were approximately the same length. Thus, we processed both channels using the same radius.
Figure 4. Background subtraction on SV2 and α-BTX channels. A, B. Images of SV2- and α-BTX-labeled objects before background subtraction. C, D. Images of SV2- and α-BTX-labeled objects after correcting the uneven background illumination using the rolling ball background subtraction method.
3. Image segmentation:
The aim of this step is to separate the objects or particles (foreground) from the background by global thresholding using pixel intensity.
a. Begin by splitting the SV2 and α-BTX channels from the image. To split channels, choose Image → Color → Split channels.
Note: You can threshold without splitting the channels. However, it is recommended to split the channels into different image windows.
b. Then, choose Image → Adjust → Threshold. A Threshold dialog box will appear. Select the α-BTX channel and click on the Set option on the Threshold dialog box to enter a cutoff value for thresholding (indicated by the red arrow in Figure 5).
c. Enter the Lower threshold level as 45 (as indicated by the black arrow in Figure 5) on the Set Threshold Levels dialog box and click OK. This step will divide the image into two classes of pixels: pixels greater than the cutoff value (foreground in red) and pixels less than the cutoff value (background in black).
Note: You can invert colors for foreground and background on the Threshold dialog box.
d. Click Apply on the Threshold dialog box to convert the image to a binary image.
e. Repeat steps D3b–d on the SV2 channel to similarly create a binary image for the SV2 channel.
Note: The thresholding can be carried out using manual or auto-thresholding options. An auto threshold is often considered more reproducible and unbiased. However, thresholding works on the basis of distributing pixel intensities on the images. If the distribution changes, thresholding will change too. Thus, even auto threshold can introduce errors by changing the thresholding cutoff levels. In practice, one should determine the method and cutoff value for thresholding using positive and negative controls. Count the number of objects in one or more control images (in a blinded manner) and then apply different thresholding methods to compare the object count with the control images. Determine the method that works best for your experiments.
Figure 5. Image segmentation by using pixel intensity thresholding. Here, the objects are represented in red on a black background. The red arrow on the Threshold dialog box on the right shows the Set option. Clicking the Set option opens the Set Threshold Levels dialog box. The black arrow on the Set Threshold Levels dialog box shows the value for the lower threshold level.
4. Apply the watershed plugin on the binary images to separate out any overlapping particles. Select the binary image for α-BTX channel and choose Process → Binary → Watershed from the ImageJ menu. Repeat this step for the binary image generated for the SV2 channel. As illustrated in Figure 6, the algorithm will separate out the connected objects (compare the regions pointed out by the arrows in panels C and F).
Figure 6. Watershed method for separating two or more connected objects. A, B. Binary images of SV2 and α-BTX channels generated after the image thresholding in step D3. D, E. Watershed images of A and B after applying the watershed plugin. C, F. Insets from B and E. The arrow in F indicates the objects separated by the watershed method.
5. Counting particles:
In this step, use the Analyze Particles tool to count the number of objects or particles in the segmented images of α-BTX and SV2 channels.
a. Click on the segmented image generated after step D4 and choose Analyze → Analyze Particles. The Analyze Particles dialog box appears. This tool allows us to filter out particles of specific size and circularity range.
b. To count the number of SV2- and α-BTX-labeled particles with an area greater than 8 µm2, input 8-Infinity for Size (micron^2) on the Analyze Particles dialog box as illustrated in Figure 7. We determined the 8 µm2 value after trying out multiple values. At this cutoff, we were able to eliminate most particles that were visibly outside the muscles and thus were not NMJs.
c. NMJs are not necessarily circular, so leave the Circularity as 0.00–1.00 (value 0 denotes a line, and value 1 indicates a perfect round object). Thus, this option will allow us to select particles of all shapes.
d. Choose Masks from the Show dropdown menu and check the boxes Summarize and Overlay on the Analyze particles dialog box. Click OK to find the object count on a result table.
e. Record the particle count for SV2- and α-BTX-labeled objects for all untreated and ethanol-exposed zebrafish samples. Figure 8 shows representative images after the area selection, segmentation, and analyze particles methods.
Note 1: A macro can be recorded on ImageJ using the Macro recorder to automate these set of commands. Alternatively, repeat these steps manually for both SV2 and α-BTX channels on untreated and ethanol-exposed zebrafish images, keeping all parameters of image analyses and image acquisition the same across all images.
Note 2. This method has been used here to separately analyze presynaptic and postsynaptic particles. However, the protocol can be modified to count the number of NMJs by looking at the co-localized SV2 and α-BTX particles using the AND function on Image Calculator. After correcting the background illumination in step D2 of this protocol, split the SV2 and α-BTX channels from the image. To split channels, choose Image → Color → Split Channels. Then, go to Image Calculator by selecting Process → Image Calculator. An Image Calculator dialog box will appear. Choose SV2 channel as Image1 and α-BTX channel as Image2. Select AND as the operation, check on the Create new window box, and click OK. A new image will be generated with only co-localized SV2 and α-BTX particles. Segment this image and analyze particles (using steps D3, 4, and 5) to get the NMJ count.
Figure 7. Counting particles using the Analyze Particles tool on ImageJ. Define the size and circularity range on the Analyze Particles dialog box. Here, particles greater than 8 µm2 were filtered out using this tool.
Figure 8. Representative images generated after applying the area selection, segmentation, and particle count methods for SV2- and α-BTX-labeled objects in untreated and alcohol-treated zebrafish. A–D. Cropped region of interest with an area of 300 × 180 µm2 after applying the area selection tool to untreated (A, B) and ethanol-treated (C, D) images. E-H. Segmented images with objects in white on a black background. Binary images of untreated (E, F) and ethanol-treated (G, H) SV2 and α-BTX channels obtained after manual thresholding using a specific cutoff value of 45 as the lower threshold level. I–L. Selected objects with size greater than 8 µm2 included in the final count using the Analyze Particles tool in untreated (I, J) and ethanol-treated (K, L) SV2 and α-BTX images.
Data analysis
Plot the number of recorded particles, presynaptic terminals (SV2-labeled particles), and postsynaptic terminals (α-BTX-labeled particles) in the control and alcohol-exposed zebrafish samples using GraphPad Prism software. Enter the data in a Grouped table format and run a two-way ANOVA with Tukey’s multiple comparisons correction to compare the number of presynaptic and postsynaptic terminals in the untreated and ethanol-exposed zebrafish. Present the data as mean ± SEM. We have previously used this method with n = 14 for each treatment. The detailed description can be found in section 4.10 and Figure 11S of Ghosal et al., 2023. Using this protocol, we found a reduction in the number of postsynaptic terminals relative to presynaptic terminals in alcohol-treated zebrafish.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article:
Ghosal et al. [9]. Embryonic ethanol exposure disrupts craniofacial neuromuscular integration in zebrafish larvae. Front Physiol (Figure 11, panel G–S).
Acknowledgments
This work was funded by NIH/NIAAA R01AA023426, NIH/NIDCR R35DE029086, and NIH/NIAAA R01AA031346 to JKE. This protocol was originally described and validated by Ghosal et al. [9].
Competing interests
The authors declare no competing interests. This work was conducted in the absence of any commercial or financial relationships.
Ethical considerations
The zebrafish lines used for this study were housed and raised in the University of Texas at Austin based on previously described protocols [13]. All protocols were approved by the Institutional Animal Care and Use Committee.
References
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Jing, L., Gordon, L. R., Shtibin, E. and Granato, M. (2010). Temporal and Spatial Requirements of unplugged/MuSK Function during Zebrafish Neuromuscular Development. PLoS One. 5(1): e8843. https://doi.org/10.1371/journal.pone.0008843
Singh, J. and Patten, S. A. (2022). Modeling neuromuscular diseases in zebrafish. Front Mol Neurosci. 15: e1054573. https://doi.org/10.3389/fnmol.2022.1054573
Singh, J., Pan, Y. E. and Patten, S. A. (2023). NMJ Analyser: a novel method to quantify neuromuscular junction morphology in zebrafish. Bioinformatics. 39(12): e1093. https://doi.org/10.1093/bioinformatics/btad720
Lescouzères, L., Bordignon, B. and Bomont, P. (2022). Development of a high-throughput tailored imaging method in zebrafish to understand and treat neuromuscular diseases. Front Mol Neurosci. 15: e956582. https://doi.org/10.3389/fnmol.2022.956582
Luderman, L. N., Michaels, M. T., Levic, D. S. and Knapik, E. W. (2022). Zebrafish Erc1b mediates motor innervation and organization of craniofacial muscles in control of jaw movement. Dev Dyn. 252(1): 104–123. https://doi.org/10.1002/dvdy.511
Nijhof, B., Castells-Nobau, A., Wolf, L., Scheffer-de Gooyert, J. M., Monedero, I., Torroja, L., Coromina, L., van der Laak, J. A. and Schenck, A. (2016). A New Fiji-Based Algorithm That Systematically Quantifies Nine Synaptic Parameters Provides Insights into Drosophila NMJ Morphometry. PLoS Comput Biol. 12(3): e1004823. https://doi.org/10.1371/journal.pcbi.1004823
Schindelin, J., Arganda-Carreras, I., Frise, E., Kaynig, V., Longair, M., Pietzsch, T., Preibisch, S., Rueden, C., Saalfeld, S., Schmid, B. et al. (2012). Fiji: an open-source platform for biological-image analysis. Nat Methods. 9(7): 676–82. https://doi.org/10.1038/nmeth.2019
Ghosal, R., Borrego-Soto, G. and Eberhart, J. K. (2023). Embryonic ethanol exposure disrupts craniofacial neuromuscular integration in zebrafish larvae. Front Physiol. 14: e1131075. https://doi.org/10.3389/fphys.2023.1131075
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Article Information
Publication history
Received: Oct 15, 2024
Accepted: Jan 2, 2025
Available online: Jan 26, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Category
Developmental Biology > Morphogenesis
Developmental Biology
Biological Sciences > Biological techniques
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Rapid and Efficient Isolation of Total RNA-Bound Proteomes by Liquid Emulsion–Assisted Purification of RNA-Bound Protein (LEAP-RBP)
JK JohnCarlo Kristofich
CN Christopher V. Nicchitta
Published: Vol 14, Iss 14, Jul 20, 2024
DOI: 10.21769/BioProtoc.5236 Views: 841
Reviewed by: Dipak Kumar PoriaThirupugal Govindarajan Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Nature Communications Sep 2023
Abstract
The critical roles of RNA-binding proteins (RBPs) in all aspects of RNA biology fostered the development of methods utilizing ultraviolet (UV) crosslinking and method-specific RNA enrichment steps for proteome-wide identification and assessment of RBP function. Despite the substantial contributions of these UV-based RNA-centric methods to our understanding of RNA–protein interaction networks, their utility is constrained by biases in RBP recovery and significant noise contributions, which can confound meaningful interpretation. To overcome these issues, we recently developed a method termed Liquid Emulsion–Assisted Purification of RNA-Bound Protein (LEAP-RBP) and introduced quantitative signal-to-noise (S:N)-based metrics for the proteome-wide identification of RNA interactomes and accurate assessment of global RBP occupancy dynamics. Compared to existing methodologies, LEAP-RBP provides significant advantages in speed, cost, efficiency, and selectivity for RNA-bound proteins. In this work, we provide a step-by-step guide for the successful application of the LEAP-RBP method for both small- and large-scale investigations of RNA-bound proteomes.
Key features
• Unbiased and efficient isolation of total RNA-bound protein, RNA, and protein from biological samples.
• Cost-effective identification of proteome-wide RNA interactomes and validation of direct RNA-binding protein functionality.
• Robust and accurate assessment of context- and/or condition-dependent RBP occupancy state dynamics.
Keywords: LEAP-RBP RNA-binding protein UV crosslinking RNA interactome RNA binding protein profiling
Graphical overview
Background
With the growing interest in the identification and study of RNA interactomes, an expanding toolkit of methods to capture RNA-protein complexes has appeared [1–9]. The most widely utilized methods employ UV crosslinking and different RNA-enrichment or organic phase–separation conditions for selective isolation of RNA–protein complexes and can be categorized as UV-based RNA-centric methods, or RNA-centric methods for short [10–17]. When paired with downstream quantitative mass spectrometry, these methods address the need for proteome-wide identification of candidate RNA-binding proteins (RBPs). We recently reported an RNA-centric method termed Liquid Emulsion–Assisted Purification of RNA-Bound Protein (LEAP-RBP), which can be distinguished from existing methods by its high selectivity for protein–RNA complexes via the use of a lithium-supplemented heterogeneous solvent system [18]. Although the precise biochemical/biophysical mechanism for the high selectivity of LEAP-RBP for protein–RNA adducts remains conjecture, we postulate that lithium complexation with the RNA component of protein–RNA adducts promotes selective precipitation, consistent with the established, but not well understood, capacity for lithium to precipitate RNA [19]. Importantly, the high selectivity of the LEAP-RBP method for protein–RNA complexes enables signal-to-noise (S:N)-based metrics for quantitative evaluation of protein–RNA interactions. Combined, LEAP-RBP addresses limitations in existing methodologies, which include biases in RBP recovery due to the inclusion of stringent denaturing washes, confounding levels of experimental noise (i.e., free protein recovery in the RBP fraction), and a paucity of quantitative metrics for assessing RNA binding function [18]. The utility of the LEAP-RBP method has been further elaborated in the identification of RBPs undergoing a condition-dependent change in RNA occupancy following short-term inhibition of translation initiation [18]. These included both RBPs with well-established roles in translation-related processes such as UPF1 and RPS3 and proteins with previously unknown roles in RNA regulation such as ABCF3 [18]. In summary, the high protein–RNA complex specificity and efficiency of the LEAP-RBP method supports rapid and inexpensive orthogonal validation of RBP occupancy state changes and RNA-binding function of candidate RBPs, using basic laboratory techniques such as SDS-PAGE RNase-dependent Assay (SRA). The simplicity and broad applicability of the LEAP-RBP method position it as a valuable tool for low- and high-throughput identification of RNA-binding proteins and rigorous assessment of RBP occupancy state change dynamics.
Materials and reagents
2-mercaptoethanol (Sigma-Aldrich, catalog number: M7522)
Acidic phenol pH 4.5 (VWR, catalog number: 0981)
Chloroform, ethanol stabilizer (Thermo Fisher Scientific, catalog number: MCX10601)
Isopropanol (VWR, catalog number: BDH1133)
Methanol (MeOH) (VWR, catalog number: BDH1135)
Sodium citrate (VWR, catalog number: BDH9288)
Citric acid (Sigma-Aldrich, catalog number: C0759)
EDTA disodium salt dihydrate (VWR, catalog number: 0105)
N-lauroylsarcosine sodium salt (Sigma-Aldrich, catalog number: L9150)
Guanidine thiocyanate (Chem-Impex, catalog number: 00522)
Diethyl pyrocarbonate (DEPC) (RPI, catalog number: D43060)
Hydrochloric acid (Ricca Chemical, catalog number: RABH0010)
TRIS base (VWR, catalog number: 97062)
LiDS (Thermo Fisher Scientific, catalog number: J32816)
LiCl (VWR, catalog number: ALFA10515; Sigma-Aldrich, catalog number: 213233)
Sodium hydroxide (VWR, catalog number: BDH9292)
Turbo DNase kit (Thermo Fisher Scientific, catalog number: AM2238)
Pierce BCA kit (Thermo Fisher Scientific, catalog number: 23225)
Dulbecco’s phosphate-buffered saline (PBS) (Thermo Fisher Scientific, catalog number: 14190144)
Solutions
DEPC-treated ultrapure H2O (see Recipes)
5 M GT (see Recipes)
750 mM sodium citrate pH 7.0 (see Recipes)
10% N-lauroylsarcosine (m/v) (see Recipes)
0.5 M EDTA pH 8.0 (see Recipes)
GT buffer (see Recipes)
AGP (see Recipes)
1.0 M Tris-HCl pH 8.0 (see Recipes)
10% LiDS (m/v) (see Recipes)
1% LiDS TE (see Recipes)
2% LiDS TE (see Recipes)
10 M LiCl (see Recipes)
Precipitation solution (see Recipes)
95% MeOH (see Recipes)
Recipes
DEPC-treated ultrapure H2O (1 L)
Reagent Final concentration Amount or volume
Ultrapure H2O n/a 999.0 mL
Diethyl pyrocarbonate (DEPC) 0.1% v/v 1 mL
Total n/a 1 L
Prepare in an airtight glass container. Shake vigorously for 5 s to dissolve DEPC. Incubate overnight at 37 °C and autoclave to decompose unreacted DEPC. Shelf life is undetermined, > 1 year.
5 M GT (0.5 L)
Reagent Final concentration Amount or volume
Guanidine thiocyanate 5 M 295.4 g
DEPC-treated ultrapure H2O n/a 271 mL
Total n/a 0.5 L
Add in the order listed. Weigh guanidine thiocyanate and adjust the volume of DEPC-treated ultrapure H2O accordingly. Prepare in a beaker using a magnetic stir bar and hot plate stirrer set to 37 °C. Clarify by filtering twice using Whatman paper or by standing incubation overnight and transferring the clarified portion. Store in an airtight glass container at room temperature (RT). Shelf life is undetermined, > 1 year. Protect from light during all stages of the preparation procedure.
750 mM sodium citrate pH 7.0 (50 mL)
Reagent Final concentration Amount or volume
Citric acid anhydrous n/a 0.13 g
Sodium citrate dihydrate n/a 10.83 g
DEPC-treated ultrapure H2O n/a Adjust volume to 50 mL
Total n/a 50 mL
Add in the order listed. Prepare in a 50 mL conical tube. Weigh citric acid anhydrous and adjust the amount or volume of other components accordingly. Dissolve by incubating on a rotator at RT. Filter using a syringe equipped with a 0.2 µm cellulose acetate syringe filter. Store at RT. Shelf life is undetermined, > 1 year.
10% N-lauroylsarcosine m/v (10 mL)
Reagent Final concentration Amount or volume
N-lauroylsarcosine sodium salt 10% m/v 1 g
DEPC-treated ultrapure H2O n/a 9 mL
Total n/a 10 mL
Add in the order listed. Prepare in a 15 mL conical tube. Weigh N-lauroylsarcosine sodium salt and adjust the volume of DEPC-treated ultrapure H2O accordingly. Dissolve by incubating on a rotator at RT. Store at 4 °C. Shelf life is undetermined, > 6 months.
0.5 M EDTA pH 8.0 (50 mL)
Reagent Final concentration Amount or volume
Sodium hydroxide pellets n/a 1.28 g
DEPC-treated ultrapure H2O n/a 44.27 mL
EDTA disodium salt dihydrate 0.5 M 9.31 g
Total n/a 50 mL
Add in the order listed. Prepare in a 50 mL conical tube. Weigh sodium hydroxide pellets and adjust the amount or volume of other components accordingly. Dissolve by incubating on a rotator at RT. Filter using a syringe equipped with a 0.2 µm cellulose acetate syringe filter. Store at RT. Shelf life is undetermined, > 1 year.
GT buffer (50 mL)
Reagent Final concentration Amount or volume
DEPC-treated ultrapure H2O n/a 5 mL
5 M GT 4 M 40 mL
750 mM sodium citrate pH 7.0 25 mM 1.67 mL
0.5 M EDTA pH 8.0 5 mM 500 µL
10% N-lauroylsarcosine m/v 0.5% m/v 2.5 mL
2-mercaptoethanol 0.7% v/v 350 µL
Total n/a 50 mL
Add in the order listed. Prepare in a 50 mL conical tube. Can be prepared without N-lauroylsarcosine and 2-mercaptoethanol and stored at RT. Protect from light. Shelf life of GT buffer (−N-lauroylsarcosine m/v and 2-mercaptoethanol) is undetermined, > 1 year. Use within a day after adding of N-lauroylsarcosine and 2-mercaptoethanol or store at -80 °C (shelf life is undetermined, > 1 year). Mix stock solutions before adding.
AGP (10 mL)
Reagent Final concentration Amount
GT buffer 66% v/v 6.67 mL
Acidic phenol pH 4.5 n/a 3.33 ml
Total n/a 10 mL
Add in the order listed. Combine reagents in a 15 mL conical tube and store at RT. Protect from light. Use within a day or store at -80 °C (shelf life is undetermined, > 1 year).
1.0 M Tris-HCl pH 8.0 (50 mL)
Reagent Final concentration Amount or volume
TRIS base 1.0 M 6.06 g
DEPC-treated ultrapure H2O n/a 43.1 mL
Hydrochloric acid n/a 2.35 mL
Total n/a 50 mL
Add in the order listed. Prepare in a 50 mL conical tube. Dissolve by incubating on a rotator at RT. Filter using a syringe equipped with a 0.2 µm cellulose acetate syringe filter. Store at RT. Shelf life is undetermined, > 1 year.
10% LiDS m/v (50 mL)
Reagent Final concentration Amount or volume
LiDS 10% m/v 5 g
DEPC-treated ultrapure H2O n/a Adjust volume to 50 mL
Total n/a 50 mL
Add in the order listed. Prepare in a 50 mL conical tube. Weigh LiDS and adjust the volume of DEPC-treated ultrapure H2O accordingly. Dissolve by incubating on a rotator at RT. Store at 4 °C. Shelf life is undetermined, > 6 months.
1% LiDS TE (50 mL)
Reagent Final concentration Amount or volume
DEPC-treated ultrapure H2O n/a 44.4 mL
1 M Tris-HCl pH 8.0 10 mM 500 µL
0.5 M EDTA pH 8.0 1 mM 100 µL
10% LiDS m/v 1% m/v 5 mL
Total n/a 50 mL
Add in the order listed. Prepare in a 50 mL conical tube and store at RT. Shelf life is undetermined, > 1 year.
2% LiDS TE (2 mL)
Reagent Final concentration Amount or volume
DEPC-treated ultrapure H2O n/a 1,552 µL
1 M Tris-HCl pH 8.0 20 mM 40 µL
0.5 M EDTA pH 8.0 2 mM 8 µL
10% LiDS m/v 2% m/v 400 µL
Total n/a 2 mL
Add in the order listed. Prepare in a 2 mL microcentrifuge tube and store at RT. Shelf life is undetermined, > 1 year.
10 M LiCl (50 mL)
Reagent Final concentration Amount or volume
LiCl 10 M 21.20 g
DEPC-treated ultrapure H2O n/a 39.76 mL
Total n/a 50 mL
Add in the order listed. Prepare in a 50 mL conical tube. Add LiCl quickly and close the cap. Adjust the volume of DEPC-treated ultrapure H2O accordingly to limit moisture absorption. Add water to LiCl salt while on ice. Close tube and incubate on ice with occasional inversion. Finish dissolving LiCl by incubating on a rotator at RT. Avoid exposing the tube seal to LiCl salt to prevent leakage. Clarify by centrifugation at 3,000–3,200× g for 30 min at RT, transfer 90% v/v of the clarified portion to a new 50 mL conical tube, and store at RT. Shelf life is undetermined, > 1 year.
Precipitation solution (40 mL)
Reagent Final concentration Amount or volume
DEPC-treated ultrapure H2O n/a 5 mL
10 M LiCl 3.75 M 15 mL
100% isopropanol v/v 50% v/v 20 mL
Total n/a 40 mL
Add reagent components to DEPC-treated ultrapure H2O. Prepare in a 50 mL conical tube and store at RT. Shelf life is undetermined, > 1 year. A small decrease in volume to 36–38 mL after mixing components is expected.
95% MeOH (1 L)
Reagent Final concentration Amount or volume
100% MeOH v/v 95% v/v 950 mL
DEPC-treated ultrapure H2O n/a 50 mL
Total n/a 1 L
Add in the order listed. Store in a glass airtight bottle and/or aliquot into 50 mL conical tubes. Store in flammable storage cabinet when not in use. Shelf life is undetermined, > 1 year. Alternatively, add 210.5 mL of DEPC-treated ultrapure H2O to an unopened 4 L bottle of MeOH.
Laboratory supplies
15 cm culture plates (e.g., Corning, catalog number: 430599)
Cell lifter (e.g., BIOLOGIX, catalog number: 70-2180)
30 mL syringe (e.g., VWR, catalog number: 76124-668)
18–22 G hypodermic needles (e.g., BD, catalog number: 305187)
Sterile syringe filter w/ 0.2 µm cellulose acetate membrane (e.g., VWR, catalog number: 28145-477)
1.5 and 2.0 mL microcentrifuge tubes (1.5 mL tubes must have round bottoms) (e.g., VWR, catalog numbers: 490004-444, 525-1136)
0.2 mL thermocycler tubes (e.g., VWR, catalog number: 20170-012)
15 and 50 mL conical tubes (e.g., Corning, catalog numbers: 430790, 430828)
0.5 and 1 L airtight glass bottles (e.g., VWR, catalog numbers: 89000-238, 89000-240)
Whatman paper (e.g., Sigma-Aldrich, catalog number: WHA1001150)
10, 200, and 1 mL pipette tips (e.g., Genesee Scientific, catalog numbers: 24-121RL, 24-150RS, 23-165RL)
10, 100, 200, and 1,000 µL pipettes (e.g., Gilson or Eppendorf)
10 and 25 mL serological pipettes (e.g., Genesee Scientific, catalog numbers: 12-104, 12-106)
Pipette controller (e.g., Thermo Fisher Scientific, catalog number: 14-387-165)
Kimwipes (e.g., VWR, catalog number: 82003-820)
96-well microplate (e.g., VWR, catalog number: 82050-678)
Thermal adhesive sealing film (e.g., Thermo Fisher Scientific, catalog number: 08-408-240)
Nitrile gloves (e.g., Microflex, catalog number: 92-134)
Repeater pipette (e.g., Eppendorf, catalog number: 022260201)
5 mL Combitips (e.g., Sigma-Aldrich, catalog number: EP0030089456)
Equipment
UV crosslinker (e.g., Stratagene, model: Stratalinker UV 2400 Crosslinker)
Analytical balance (e.g., Mettler Toledo, model: ME54E)
Sample tube rotator (e.g., Fisher Scientific Hematology Chemistry Mixer, model: 346)
Tube rack (e.g., VWR, catalog number: 82010-750)
Refrigerated microcentrifuge (e.g., Eppendorf, model: 5417R)
Centrifuge with 15 mL and 50 mL conical tube holders (e.g., Beckman, model: CS-6R)
Thermocycler (e.g., Bio-Rad, model: T100 PCR Thermocycler)
Hot plate stirrer (e.g., Accuplate, model: D0310)
UV spectrophotometer (e.g., NanoDrop, model: ND-100)
Microplate reader (e.g., VersaMax, model: M5e)
Dry bath/heating block (e.g., Thermolyne, model: 17600)
Mini centrifuge (e.g., ISC BioExpress, model: C1301P-ISC)
37 °C incubator (e.g., Baxter Scientific Products, model: J1450-3)
4 °C refrigerator (e.g., Kenmore, model: 253.6880201E)
-20 °C freezer (e.g., Kelvinator, model: KFU21M7LW3)
-80 °C freezer (e.g., VWR, model: 5706)
Software and datasets
Microsoft Excel for Mac version 16.66.1
SoftMax Pro version 7.2
NanoDrop 1000 version 3.8.1
Procedure
Cell culturing, UV crosslinking, and cell harvest
Grow cells to the desired confluency in a 15 cm plate.
Note: Adjust volumes used during subsequent steps accordingly when employing plates of varying sizes.
Decant media and place the cell culture dish on an ice-filled culture dish lid.
Wash cells twice with 12 mL of ice-cold 1× PBS.
Tilt the plate to an 80° angle while on the ice-filled culture dish lid, incubate for 1 min, and aspirate residual PBS.
Irradiate cells while on the ice-filled culture dish lid with the desired amount of UV energy and at the desired wavelength (e.g., 0.4 J/cm2, 254 nm) using a Stratalinker or other suitable UV crosslinker (General note 1).
Lyse, scrape, and transfer cells into a 2.0 mL microcentrifuge tube on ice using two 0.4 mL aliquots of GT buffer and a cell scraper.
Critical: Pre-wet pipette tips for pipetting accuracy (General note 2).
Add 0.4 mL of acidic phenol for a total of 1.2 mL of AGP (i.e., 0.8 mL of GT and 0.4 mL of acidic phenol) and sheer the lysate by passaging through a 19–20 G needle 15 times. Add additional AGP to a final concentration of 80% v/v.
Note: Keep samples at RT after sheering unless indicated otherwise. Sheered lysates represent starting AGP input suspensions (Table S1).
Caution: Perform this step in a fume hood.
Caution: Avoid exposing tube seals to AGP to prevent sample leakage (General note 3).
Critical: Pre-wet pipette tips for pipetting accuracy (General note 2).
Critical: Mix stock solutions and sample suspensions containing AGP < 10 s prior to sampling (General note 4).
Pause point: Samples can be stored at RT in AGP for 16 h or at -80 °C for long-term storage (General note 5).
For initial compositional analysis of starting AGP input suspensions, transfer two 10–50 µL aliquots to separate 1.5 mL microcentrifuge tubes containing an appropriate volume of AGP for a final volume of 200 µL. Isolate RNP fractions according to the LEAP-RBP method (steps C1–C9), suspend in 10–50 µL of 1% LiDS TE as described in step F1, and perform RNA (UV spectrophotometry) quantitation as described in step F3. Estimate RNA and protein yields of the starting AGP input suspension using the tools included as part of Table S1.
Caution: Perform dilution of starting AGP input suspensions in fume hood.
Caution: Avoid exposing tube seals to AGP to prevent sample leakage (General note 3).
Critical: Pre-wet pipette tips for pipetting accuracy (General note 2).
Critical: Mix stock solutions and sample suspensions containing AGP < 10 s prior to sampling (General note 4).
Pause point: Samples can be stored at RT for 16 h or at -80 °C for long-term storage (General Note 5).
Add AGP to the undiluted starting AGP input suspension for a final concentration of 10–250 ng of RNA and DNA/µL and > 86% AGP v/v for isolation of RNP fractions by LEAP-RBP or 25–2,000 ng of protein/µL for isolation of total input samples by 95% MeOH v/v precipitation (Sections C, E).
Note: A dilution calculator is included as part of Table S1.
Caution: Perform this step in a fume hood.
Caution: Avoid exposing tube seals to AGP to prevent sample leakage (General note 3).
Critical: Pre-wet pipette tips for pipetting accuracy (General note 2).
Critical: Mix stock solutions and sample suspensions containing AGP < 10 s prior to sampling (General note 4).
Pause point: Samples can be stored at RT for 16 h or at -80 °C for long-term storage (General note 5).
Repeated AGPC extraction
Transfer 1 mL aliquots of an AGP input suspension containing > 80% AGP v/v to separate 2.0 mL microcentrifuge tubes (General note 8). Add 200 µL of chloroform per milliliter of AGP (v/v), vortex (10 s, maximum setting), and centrifuge at 10,000× g for 10 min at 4 °C with slow brake setting. Carefully remove 80%–90% of the aqueous and organic phases without removing the AGPC interphase (General note 6).
Note: Slow or “soft” brake settings can be found on many tabletop centrifuges (e.g., Eppendorf, model: 5417R). By slowing the rate of deceleration, the likelihood of disturbing the pellet fraction is reduced. Not including this setting when listed may impact yield and/or introduce biases, although this has not yet been established.
Caution: Perform this step in a fume hood.
Caution: Avoid exposing tube seals to AGP or AGPC to prevent sample leakage (General note 3).
Critical: Pre-wet pipette tips for pipetting accuracy (General note 2).
Critical: Mix stock solutions and sample suspensions containing AGP < 10 s prior to sampling (General note 4).
Perform additional AGPC extractions as described in the previous step using 800 µL of AGP and 160 µL of chloroform per 2.0 mL microcentrifuge tube per extraction.
Note: AGPC interphase samples can be pooled 2:1 before each additional AGPC extraction (General note 7). The required number of repeated AGPC extractions depends on protein UV-crosslinking efficiency and sample composition (General note 9).
Caution: Perform this step in a fume hood.
Caution: Avoid exposing tube seals to AGP or AGPC to prevent sample leakage (General note 3).
Critical: Remove most of the organic phase before the next step.
Critical: Pre-wet pipette tips for pipetting accuracy (General note 2).
Critical: Mix stock solutions and sample suspensions containing AGP < 10 s prior to sampling (General note 4).
Add 1 mL of AGP to each final AGPC interphase sample and pool if applicable. Determine RNA yield according to step A8. Dilute final AGPC interphase suspensions to 10–250 ng of RNA/µL for isolation of RNP fractions by LEAP-RBP (Section C).
Caution: Perform this step in a fume hood.
Caution: Avoid exposing tube seals to AGP or AGPC to prevent sample leakage (General note 3).
Critical: Pre-wet pipette tips for pipetting accuracy (General note 2).
Critical: Mix stock solutions and sample suspensions containing AGP < 10 s prior to sampling (General note 4).
Pause point: Samples can be stored at RT for 16 h or at -80 °C for long-term storage (General note 5).
Isolation of RNP fractions by LEAP-RBP
Transfer 200 µL aliquots of an AGP input suspension or final AGPC interphase suspension to separate 1.5 mL microcentrifuge tubes (General note 8). Mix by continuous vortex within 1 h of the next step (10 s, setting 4–5). Keep samples off the lid.
Caution: Perform this step in a fume hood.
Caution: Avoid exposing tube seals to AGP or AGPC to prevent sample leakage (General note 3).
Critical: Pre-wet pipette tips for pipetting accuracy (General note 2).
Critical: Mix stock solutions and sample suspensions containing AGP < 10 s prior to sampling (General note 4).
Pause point: Samples can be stored at RT for 16 h or at -80 °C for long-term storage (General note 5).
To each 1.5 mL microcentrifuge tube: Add 14 µL of chloroform and mix by pulse vortex until an emulsion forms (1 s intervals, setting 4–5). Then, mix by continuous vortex (10 s, setting 4–5).
Note: Final AGPC interphase suspensions contain residual chloroform. Use a single 200 µL aliquot to determine the optimal amount of chloroform for the remaining aliquots. Briefly, add 12 µL of chloroform and mix by pulse vortex (1 s intervals, setting 4–5). If an emulsion does not form and/or if the emulsion takes on a more greyish appearance (see Kristofich and Nicchitta [18], Supplementary Figure 11c), add an additional 2 µL of chloroform and use 14 µL of chloroform for each of the remaining 200 µL aliquots. Otherwise, use 12 µL of chloroform.
Caution: Perform this step in a fume hood.
Caution: Avoid exposing tube seals to AGP or AGPC to prevent sample leakage (General note 3).
Critical: Pre-wet pipette tips for pipetting accuracy (General note 2).
Critical: Close sample tubes within 10 min of adding chloroform and vortex within 1 h to avoid a decrease in yield (General note 10).
If samples were stored at -80 °C and/or if starting from Turbo DNase digests, mix samples by vortex before adding chloroform (10 s, setting 4–5).
Gently add/layer 856 µL of RT precipitation solution (3.75 M LiCl, 50% isopropanol v/v) on top of each AGPC mixture and close the tubes.
Note: Add the first 100–200 µL slowly to the inner side of the sample tubes below the seal line.
Caution: Perform this step in a fume hood.
Caution: Avoid exposing tube seals to AGP, AGPC, or precipitation solution to prevent sample leakage (General note 3). Note that water condensation near the seal of the tube, which forms above the precipitation solution, will not cause sample leakage during subsequent steps.
Critical: Pre-wet pipette tips for pipetting accuracy (General note 2).
Critical: Perform this step within 1 h of the previous step to avoid a decrease in yield (General note 10).
Using a sample tube rack, gently invert tubes to a 90° angle (> 1 s) and hold until the AGPC mixtures become displaced from the bottom of the tubes (0.5–1.0 s). Revert the tubes back to an upright position at a slightly faster speed (0.5–1.0 s) and let stand for 1 min at RT. Repeat this process twice more in alternating directions for three rounds of gentle inversions/reversions followed by additional rounds with increasing angle and speed. Once most of the AGPC mixture becomes displaced from the bottom of vertically standing sample tubes, perform a final round inversion with a forceful reversion, let stand for 30 s, and vortex (5 s, maximum setting).
Note: Sample appearance and behavior will differ depending on clRNP and protein content of AGP suspensions. While this process can be completed after only five inversions, trying to complete the process in ten or more inversions will reduce the chance of overmixing without compromising results (Video S1).
Critical: Perform the first inversion/reversion step within 1 h of adding the precipitation solution and/or complete all inversion/reversion steps within 1 h and 10 min to avoid an increase in free protein recovery.
Pause point: Samples can be stored at RT for 16 h after mixing by vortex (General note 5).
Using a sample tube rack, perform two inversions with forceful inversions in opposite directions. Then, centrifuge sample tubes at 14,000× g for 5 min at 20 °C.
Critical: Not performing inversions with sufficient force results in scattered or non-uniform RNP pellets, which can negatively impact yields.
Pause point: Samples can be stored at RT for 16 h (General note 5).
Remove supernatants and partially close lids. Add 1 mL of RT 95% MeOH v/v to each sample tube and close lids. Complete the following process for one sample tube at a time: Invert three times, pausing briefly for < 1 s before each reversion, and quickly remove/discard the MeOH washes using a syringe equipped with an 18–22 G needle. Keep tube lids partially closed between MeOH washes. Repeat the process twice more using two instead of three inversions.
Note: Allow samples to incubate in 95% MeOH v/v for at least 5 min across all three washes. The white precipitate, which forms after adding 95% MeOH v/v, will be removed during the washes. It is not necessary to remove residual 95% MeOH v/v adhering to the cap or sides of tubes.
Caution: Perform this step in a fume hood.
Critical: Not removing the washes quickly after inverting tubes allows precipitates containing free protein to settle and contaminate RNP fractions.
Critical: If an RNP pellet is not dislodged from the bottom of the tube before removing the final MeOH wash, gently nudge the top and/or side of the pellet with a syringe or pipette tip and resume at the inversion step.
Critical: If RNP pellets are small and thus fragile, add and remove 95% MeOH v/v washes using a P1000 to avoid sample loss.
Pause point: Samples can be stored at RT in any of the three 95% MeOH v/v washes for 16 h (General note 5).
Transfer and/or pool RNP pellets into a new 1.5 mL microcentrifuge tube. Briefly, pour 1 mL of RT 95% MeOH v/v from a new tube into the tube containing the RNP pellet at a moderate rate for 0.5–1.0 s until the RNP pellet becomes displaced from the bottom of the tube and then quickly pour it back into the new tube.
Note: If pooling multiple RNP pellets, wait until RNP pellets have settled in the new tube before repeating the process.
Caution: Perform this step in a fume hood or limit exposure to MeOH.
Pause point: RNP pellets can be stored in 95% MeOH v/v at RT for 16 h or at -80 °C for long-term storage after being transferred to a new tube (General note 5).
Remove most of the 95% MeOH v/v leaving 50–200 µL. Centrifuge briefly and use a P1000 pipette equipped with a P10 pipette tip attached to a P1000 pipette tip to remove residual 95% MeOH v/v. Air dry pellets by incubating the tube with the lid open for 5–10 min at RT.
Note: RNP pellets, which have been pooled into the same tube, should be spread out while removing residual 95% MeOH v/v so they dry more uniformly.
Caution: Perform this step in a fume hood or limit exposure to MeOH.
Pause point: Opened sample tubes can be incubated overnight at RT if necessary. Dried RNP pellets can be stored at -80 °C for long-term storage (General note 5).
For RNP fractions isolated from diluted starting AGP input suspension (step A8), resuspend as described in step F1, perform UV-spectrophotometric analysis of unclarified sample suspensions as described in step F3, and analyze the data using the tools provided in Table S1. Otherwise, proceed to DNA depletion (Section D).
DNA depletion
Add 15 µL of TE buffer to dry RNP pellet(s) containing 10–50 µg of RNA and DNA and incubate for > 2 min at RT. Using the same pipette tip, pipette 5 µL of the sample suspension 8 times without submerging the pipette tip in the sample suspension. Incubate for > 2 min and repeat the pipetting step using the same pipette tip. During the second pipetting step, scrap the bottom of the tube while ejecting.
Note: Pooling RNP pellets to maximize the amount of RNA and DNA per tube reduces the time-sensitivity of the second LEAP step following Turbo DNase digestion (General note 10). An empty pipette tip box is a convenient way to store pipette tips between pipetting steps.
Critical: Not maintaining sample suspensions at the bottom of the tube during the resuspension procedure results in scattered precipitates during the second LEAP step and lower yields.
Critical: If dried RNP pellets have become dislodged from the bottom of the tube prior to adding TE buffer, centrifuge sample tubes at 3,000× g for 5 s.
Critical: Submerging the pipette tip and/or scraping the bottom of the tube during the first pipetting step can result in hydrated RNP pellets becoming stuck to the side of the pipette tip. In these scenarios, gently remove the pipette tip and allow it to incubate while submerged in the sample suspension between pipetting steps. Do not attempt to rub the pipette tip against the sides of the tube.
Pause point: Sample suspensions, which were kept at the bottom of the tubes, can be stored at RT for 1 h.
Using a new pipette tip for each sample, add 5.0 µL of a pre-mixed master mix containing 2.0 µL of 10× Turbo DNase Buffer and 1.0 µL of Turbo DNase enzyme per 10.0 µg of DNA (0.2 µL minimum and 1.5 µL maximum) supplemented with TE buffer to 5 µL.
Note: The pre-mixed master mix should be prepared after the previous step. DNA quantity based on the change in RNA quantity (UV spectrophotometry) is observed after the second LEAP step when using the maximum amount of Turbo DNase.
Critical: Not maintaining sample suspensions at the bottom of the tube results in scattered precipitates during the second LEAP step and lower yields.
Critical: UV-crosslinked samples begin to adhere to pipette tips after adding Turbo DNase buffer. Add the entire volume of the pre-mixed master mix directly to the sample suspension and swirl the pipette tip for < 3 s to mix without pipetting.
Incubate samples for 15 min at 37 °C.
Add 180 µL of AGP and mix by vortex (10 s, setting 5).
Note: Aliquots of AGP can be prepared ahead of time and stored at -80 °C (General note 5).
Caution: Perform this step in a fume hood.
Caution: Avoid exposing tube seals to AGP or AGPC to prevent sample leakage (General note 3).
Critical: Pre-wet pipette tips for pipetting accuracy (General note 2).
Critical: Perform this step within 15 min of the previous step to avoid a decrease in yield (General note 10).
Pause point: Sample tubes can be centrifuged briefly at 3,000× g (quick spin) and stored at RT for 16 h or -80 °C for long-term storage (General note 5).
Isolate RNP fractions by LEAP-RBP (start at step C2).
Isolation of input (total protein) samples
Transfer 50 µL aliquots of an AGP input suspension containing > 80% AGP v/v and 25–2,000 ng of protein/µL to separate 2 mL microcentrifuge tubes.
Caution: Perform this step in a fume hood.
Caution: Avoid exposing tube seals to AGP or AGPC to prevent sample leakage (General note 3).
Critical: Pre-wet pipette tips for pipetting accuracy (General note 2).
Critical: Mix stock solutions and sample suspensions containing AGP < 10 s prior to sampling (General note 4).
Pause point: Samples can be stored at RT for 16 h or at -80 °C for long-term storage (General note 5).
Mix samples by vortex (5 s, maximum setting), add 950 µL of RT 100% MeOH v/v, and vortex (10 s, maximum setting).
Caution: Perform this step in a fume hood.
Caution: Avoid exposing tube seals to AGP or AGPC to prevent sample leakage (General note 3).
Critical: Pre-wet pipette tips for pipetting accuracy (General note 2).
Pause point: Samples can be stored at RT for 16 h or at -80 °C for long-term storage (General note 5).
Incubate sample tubes on a rotator for 1 h at RT and centrifuge at 20,000× g for 10 min at 20 °C with slow-brake setting. Remove supernatants using a syringe equipped with an 18–22 G needle and partially close tube lids.
Caution: Perform this step in a fume hood.
Caution: Avoid exposing tube seals to AGP or AGPC to prevent sample leakage (General note 3).
Add 1 mL of RT 95% MeOH v/v to each sample tube, close lids, and vortex (5 s, maximum setting). Incubate sample tubes on a rotator for 10 min at RT, centrifuge at 20,000× g for 10 min at 20 °C with slow-brake setting, and remove supernatants using a syringe equipped with an 18–22 G needle. Repeat this process once or twice if pooling > 100 µg of protein during the next step. Keep tube lids partially closed between MeOH washes.
Note: Slow or “soft” brake settings can be found on many tabletop centrifuges (e.g., Eppendorf 5417R). By slowing the rate of deceleration, the likelihood of disturbing the pellet fraction is reduced. Not including this setting when listed may impact yield and/or introduce biases although this has not yet been established.
Caution: Perform this step in a fume hood.
Critical: Not removing the washes quickly and gently after centrifugation can result in sample loss.
Critical: Keep precipitates off the lid during centrifugation by holding sample tubes vertically and moving them in a single up-down motion. Repeat the process until white precipitates are no longer visible on the bottom of tube lids.
Pause point: Samples can be stored at RT in 95% MeOH v/v washes for 16 h (General note 5).
Use three 400 µL aliquots of RT 95% MeOH v/v to recover precipitates and transfer them into a new 1.5 mL microcentrifuge tube (General note 11). Keep tube lids partially closed after, between removing and adding 95% MeOH v/v.
Note: This process can be used to pool precipitates from multiple 2 mL microcentrifuge tubes. Additional aliquots can be used if necessary.
Caution: Perform this step in a fume hood or limit exposure to MeOH.
Pause point: Precipitates can be stored in 95% MeOH v/v at RT for 16 h or at -80 °C for long-term storage after being transferred to a new tube (General note 5).
Incubate sample tubes vertically for 30 min at RT to allow precipitates to settle at the bottom of the tube. Centrifuge at 20,000× g for 10 min at 20 °C with a slow-brake setting and remove most of the supernatant, leaving 50–200 µL. Repeat the centrifugation step and remove residual 95% MeOH v/v using a P1000 pipette equipped with a P10 pipette tip attached to a P1000 pipette tip.
Caution: Perform this step in a fume hood or limit exposure to MeOH.
Critical: Not removing the washes quickly and gently after centrifugation can result in sample loss.
Use a P20 pipette and 2 µL of RT 95% MeOH v/v to break up and spread out precipitates so they dry more uniformly. Incubate opened sample tubes for 10–30 min at RT or until precipitates are dry.
Note: Precipitates should be spread out before they start to dry.
Caution: Perform this step in a fume hood or limit exposure to MeOH.
Critical: Not spreading out precipitates will make resuspension more difficult (Section F).
Pause point: Opened sample tubes can be incubated overnight at RT if necessary. Dried precipitates can be stored at -80 °C for long-term storage (General note 5).
Sample resuspension, clarification, and quantitation
Add 1% LiDS TE to dried RNP pellets or precipitates and incubate for 30 min at RT with occasional pipetting at the 2, 15, and 30 min mark. Briefly, pipette 90% of the sample volume eight times, scraping the bottom of the tube while ejecting the sample. During the initial pipetting step (2 min mark), pipette samples until most of the visible precipitates are suspended and not adhering to the sides of the tube.
Note: The volume of 1% LiDS TE added to samples and the desired concentration depends on the intended downstream application(s). In particular, steps F2–4 are intended for samples suspended above their working concentration (i.e., 4.0 µg of RNA and DNA/µL for RNP fractions and 5.0 µg of protein/µL for input samples). If samples are suspended below their working concentrations, proceed to step F5, and adjust sample dilutions during step F6 accordingly (if applicable) so they fall within the linear quantitation range of UV spectrophotometer and/or BCA (Table S2). Efforts should be made to not introduce bubbles during the pipetting steps. Nonetheless, they will be removed during the clarification procedure. The same pipette tip can be used for all three pipetting steps by storing them in a pipette tip box between pipetting steps.
Critical: Do not store samples on ice. Samples do not contain active proteases and/or nucleases, so storage on ice is not necessary.
Critical: Samples should not be stored after starting the resuspension procedure and prior to the clarification step.
Mix samples by vortex (5 s, setting 5) and quickly (< 10 s) transfer 2 µL to a 0.2 mL thermocycler tube containing 8 µL of 1% LiDS TE for a 1:5 dilution. Rinse pipette tips after transferring by pipetting the added volume three times.
Critical: Samples suspended well above their working concentrations are difficult to pipette accurately. If this is the case, measure the pipetted sample volume using the 2 µL graduation mark that can be found on most standard P10 pipette tips.
Analyze 1–2 µL of diluted (1:5) sample suspensions by UV spectrophotometry (Nanodrop, RNA setting) and calculate the concentration of undiluted sample suspensions (Table S2).
Note: DNA and protein contributions confound RNA quantitation. Nonetheless, it provides a reliable and rapid means for diluting samples to their working concentration.
Critical: Pipette 90% of the sample volume five times < 10 s before analyzing samples.
Critical: The recommended range for quantitation of 1% LiDS TE sample suspensions is 0.1–1.8 µg of RNA&DNA for RNP fractions and 0.1–1.0 µg of RNA&DNA for input sample as determined by UV spectrophotometry (Nanodrop, RNA setting).
Critical: Read the same 1–2 µL aliquot twice. If the values vary by more than 10%, rigorously clean the pedestal of the Nanodrop UV spectrophotometer and repeat step F3. This often occurs with RNP fractions containing a high amount of crosslinked RNA-protein when they are suspended above 1.8 µg of RNA/µL and/or when not adequately mixing samples prior to UV-spectrophotometric analysis.
Add 1% LiDS TE to undiluted sample suspensions to a final concentration of 1 µg of RNA/µL for input samples and 4 µg of RNA/µL for RNP fractions as determined by UV-spectrophotometric analysis. Mix samples by brief vortex (5 s, setting 5) and incubate for 5 min at RT.
Pause point: Diluted sample suspensions can be stored at RT for 2 h.
Incubate samples in a dry bath/heating block or thermocycler at 65 °C for 3 min, vortex (5 s, maximum setting), and centrifuge at 270–310× g for 5 min at 20 °C with soft-brake setting. Carefully transfer 90% of the clarified fraction to a separate 1.5 mL microcentrifuge tube.
Note: If working with sample volumes < 100 µL, this step should be performed in 0.2 mL thermocycler tubes.
Clarified input (total protein) and RNP (total RNA-bound protein) fractions should be diluted as described in step F2 for RNA and protein quantitation.
Note: Sample dilutions can be calculated using the tools included as part of Table S2.
Pause point: Diluted and/or undiluted clarified sample suspensions can be used immediately or stored at -80 °C (General note 5).
RNA quantitation of clarified RNP fractions should be performed as described in step F3 and data should be analyzed using the tools included in Table S3 for calculation of total RNA yields.
Perform protein quantitation using the BCA microplate assay. Briefly, dilute 1 mL of a BSA standard (2 mg/mL) with 1 mL of 2% LiDS TE (D1) and use 500 µL to prepare seven 1:2 serial dilutions using 1% LiDS TE (D2–D8). Prepare the BCA working reagent and place a 96-well microplate on ice. Mix diluted clarified sample suspensions prepared in step F6 by vortex (5 s, setting 5) and quickly (< 10 s) transfer two 2–4 µL aliquots to separate wells (Table S2). Add two 2 µL aliquots of 1% LiDS TE and each of the BSA standards to separate well in descending order of their dilution number (D#). Then, add an additional two 4 µL aliquots of D1 to separate wells. Promptly (< 3 min) add 200 µL of the BSA working reagent to each well containing BSA standards in ascending order of their dilution number (D#). Lastly, add 200 µL of the BSA working reagent to each of the remaining wells containing either 1% LiDS TE or aliquoted sample suspensions. Incubate the plate on ice for 5 min, attach thermal adhesive sealing film, and incubate at 37 °C for 2.5–3 h. Remove the sealing film and measure absorbance at 562 nm using a microplate reader and appropriate software (e.g., SoftMax Pro version 7.2, standard curve setting: bi-exponential).
Note: Protein quantitation data can be exported and analyzed using the tools included in Table S3 for the determination of total protein and RNA-bound protein yields. BSA standards can be prepared ahead of time, aliquoted, and stored at -80 °C. Once thawed, BSA standards can be kept at 4 °C for 6 months. The appearance of precipitates in BSA standard D1 when stored at 4 °C is expected and will be resolved after equilibrating it to RT. Load > 0.25 µg of protein per well; non-crosslinked RNP fractions are an exception (General note 12).
Critical: Keep samples and BSA standards at room temperature when setting up the plate and use a new pipette tip for each aliquot. Add aliquots directly to the bottom side of the wells without touching the bottom, as this can scratch the plate and confound results. Add the BCA working reagent to the same side of the well.
Critical: Set up the microplate on ice to avoid the impact that differences in incubation times have on protein quantitation data.
Data analysis
Data analysis methods are detailed in the original LEAP-RBP study as part of the Supplementary Methods [18]. Briefly, LEAP-RBP fractions can be analyzed by SRA to evaluate UV-dependent enrichment and S/N of proteins. Input samples and LEAP-RBP fractions can be normalized to micrograms of protein, RNase-digested, and compared by SDS-PAGE to evaluate total and RNA-bound abundance, respectively. Total protein and total RNA crosslinking efficiencies can be estimated using RNA and protein yields of input (total protein) samples, LEAP-RBP fractions isolated from AGP input suspensions containing total RNA and total RNA-bound protein, and LEAP-RBP fractions isolated and final AGPC interphase suspensions containing total RNA-bound protein and total protein-bound RNA. SILAC LC-MS/MS analysis of LEAP-RBP fractions isolated from pooled cellular samples containing equivalent amounts of UV-crosslinked and non-crosslinked cells allows comprehensive identification of UV-enriched candidate RBPs. Comparison of input samples and corresponding LEAP-RBP fractions by LC-MS/MS informs condition- and/or context-dependent differences in total and RNA-bound abundances, respectively. Lastly, LC-MS/MS analysis of input samples and LEAP-RBP fractions allows proteome-wide determination of UV-crosslinking efficiencies (%CL) for functional validation of candidate RBPs and %CL-based ranking of protein features and classifications [20].
Validation of protocol
Validation of LEAP-RBP and the methods herein can be found in the original publication [18]. Briefly, the ability of LEAP-RBP to rapidly recover total RNA and/or bound proteins from AGP sample suspensions with near 100% efficiency was validated by SRA analysis of precipitated and unprecipitated fractions. The importance of components added during the LEAP step and the liquid–liquid interphase was evidenced by a change in RNA and/or protein yields. RNA-centricity of the LEAP-RBP method was validated by performing LEAP-RBP on RNase-treated clRNP fractions resulting in a loss of detectable protein by SDS-PAGE and Coomassie Blue staining. Efficiency of DNA depletion steps was validated by qPCR and SRA analysis. RNA isolated by LEAP-RBP method was found to be of high integrity by Bioanalyzer analysis (RIN > 9), and an RNA-seq analysis showed broad representation of diverse sRNA species displaying broad genome distributions. The efficiency of DNA and/or RNA depletion steps used for the preparation of total protein and total RNA-bound protein samples for MS proteomic was validated by compositional analyses (i.e., RNA and protein quantitation) and by comparison using both TBE gel analysis and SDS-PAGE with SYBR Safe (RNA and DNA), Coomassie Blue (protein), and Silver Stain (RNA, DNA, and protein) staining.
General notes and troubleshooting
General notes
Differences in UV-crosslinking conditions will affect total protein and protein-specific crosslinking efficiencies (Supplementary Figure 13 [18]). Importantly, this includes factors such as plate height, plate diameter, and position in the UV crosslinker. While most crosslinkers have a sensor meant to increase exposure time as the bulb energy output declines from age, it is not meant to measure the UV dose from individual bulbs and make the necessary adjustments. Because not all RNA–protein interactions exhibit similar crosslinking dose-responsiveness (Supplementary Figure 6d [18]), we highly recommend experiments to account for position-dependent differences in UV dose across experimental sample groups.
Pre-wet pipette tips by pipetting to and from a sample and/or stock solution a few times before transferring a set volume (Figure S1, Video S2).
Exposing the seal of standard laboratory tubes to AGP, AGPC, or lithium-supplemented precipitation solution (3.75 M LiCl, 50% isopropanol v/v) prior to closing the lid results in sample leakage. If this occurs, clean the seal using a Kimwipe prior to closing the tube. A snapping sound when closing the tube is indicative of a tight seal. Importantly, opening microcentrifuge tubes while AGP is on the bottom of the cap will expose gloves as evidenced by a white residue. Performing the procedures as recommended should prevent exposure and/or sample leakage. Acidic phenol poses serious health risks and can penetrate most standard laboratory gloves within minutes. Use appropriate gear and/or replace contaminated gloves under their reported penetration time. Always maintain open containers and/or sample tubes containing AGP in a fume hood.
AGP will slowly separate into two phases over time. Therefore, mix samples and/or stock solutions that contain AGP by vortex (if < 50% of tube volume: 5 s, setting 5; otherwise, 5 s, maximum setting), inversion (four times), or by pipetting (> 50% sample volume four times) < 10 s prior to sampling.
When indicated, samples can be stored at RT for a short time or at -80 °C for long-term storage (i.e., undetermined, > 1 year). We have observed negligible impact on RNA and/or protein integrity for samples even after six freeze/thaw cycles in AGP and/or 1% LiDS TE. Therefore, we recommend storing samples at -80 °C regardless of the anticipated duration. If desired, RNA integrity can be evaluated by diluting RNP fractions with TE buffer to 0.16% LiDS and analyzing 0.5 µg of RNA by TBE gel electrophoresis, while protein integrity can be evaluated by SRA analysis and total protein staining as described in the original publication [18]. Samples and/or stock solutions (e.g., aliquots of AGP) stored at -80 °C should be thawed by incubating on the bench for 30 min at RT before being used. Notably, volumes larger than 1.5 mL may require longer incubation times to thaw and equilibrate to RT.
A detailed description of the repeated AGPC extraction procedure has been included as part of the Methods and Supplementary Methods in the original publication [18]. Briefly, use a P1000 pipette tip to remove most of the aqueous phase leaving 100–150 µL. Carefully tilt the sample tube and, using a gel loading pipette tip equipped with a new P1000 pipette tip, slide down against the bottom side of the tube through a now exposed region of the organic phase until it touches the bottom of the tube. Carefully begin to aspirate the organic phase while observing the AGPC interphase and stop once it starts to break apart and/or gets close to the pipette tip. Note that this often occurs just as the AGPC interphase falls below the 250 µL graduation mark on the side of the tube. After pausing, slowly return the tube to a vertical position and attempt to remove more of the organic phase. Lastly, resume the tilted position as before and quickly remove the pipette tip at the previously used entry point.
Pooling AGPC interphase samples is an effective strategy for concentrating crosslinked RNA–protein complexes from very dilute samples. Briefly, use a new P1000 pipette to aspirate 800 µL of the AGP stock solution and eject 200–300 µL into one of two AGPC interphase samples that are going to be pooled. Continuing, aspirate the second AGPC interphase sample and transfer it to the tube containing the first AGPC interphase sample by ejecting 200–300 µL. Lastly, use the 500–600 µL of AGP remaining in the pipette tip to rinse the tube that contained the second AGPC interphase sample before transferring the entire volume back to the tube containing both AGPC interphase samples. A similar process should be used to transfer one or two AGPC interphase sample(s) to a clean 2 mL microcentrifuge tube if the seals become compromised (General note 3).
We find that accurate and consistent sampling of AGP input and final AGPC interphase suspensions requires mixing samples and pre-wetting pipette tips after every four to five aliquots using standard pipettes (General notes 2, 4). However, using a repeater pipette greatly simplifies the process and reduces the likelihood of sample leakage when sample volumes are large. For example, consider a scenario where 200 µL aliquots of a 10 mL AGP input suspension need to be transferred to separate 1.5 mL microcentrifuge tubes. If this is approached using a repeater pipette equipped with a 5 mL Combitip, the sample can be mixed by pipetting 5 mL four times before and/or after every eight 200 µL aliquots without the need for pre-wetting the pipette tip. The alternative approach using a P200 or P1000 pipette would require pre-wetting the pipette tip thereby reducing the number of aliquots that can be transferred before having to mix the sample down to four. If taking the second approach, we recommend using an appropriately sized tube to allow mixing by vortex without exposing the seal of the tube. For example, a 10 mL AGP input suspension can be mixed by vortex when it is contained in a 50 mL conical tube without it touching the tube seal (5 s, setting 5). Aliquoted samples should be centrifuged briefly (stop before getting to 3,000× g) before storing them at -80 °C (Video S1).
We recommend performing repeated AGPC extractions until the size of the interphase stops reducing in size. However, additional AGPC extractions can be employed to pool AGPC interphase samples (General note 7). The number of repeated AGPC extractions required is often between 2 and 8. Notably, additional AGPC extractions can be performed after resuspending AGPC interphase samples in AGP if desired, including those stored at -80 °C while performing an initial yield assessment (General note 5).
The time sensitivity of all steps of the LEAP-RBP has been thoroughly investigated. Briefly, we note that the time after adding chloroform and before mixing to form an AGPC mixture or after mixing to form an AGPC mixture and before adding/layering the precipitation solution can affect RNA-bound protein recovery, while the time after adding/layering the precipitation solution and before inversions/reversions can affect free protein recovery. Importantly, we did not observe an effect when limiting the duration of these steps to 1 h, and extending the duration of any step by incubating overnight at room temperature had no discernible effect on RNA integrity. More generally, we note that the time-sensitivity of steps that impact yield can be reduced by increasing the concentration of RNA and/or RNA-bound protein in AGP suspensions and the time-sensitivity of steps that impact free protein recovery can be reduced by decreasing the concentration of free protein in AGP suspensions. The most likely situation where time sensitivity becomes an issue is when performing LEAP-RBP on Turbo DNase digests containing a low amount of RNA and/or RNA-bound protein. Here, the time after adding chloroform and before mixing to form an AGPC mixture or after mixing to form an AGPC mixture and before adding/layering the precipitation solution can negatively affect yield. Therefore, we highly recommend pooling RNP pellets isolated from more diluted samples prior to the DNA depletion step and/or reducing the number of samples being processed in parallel during the time-sensitivity steps noted above.
To facilitate pooling and/or resuspension of input samples in small volumes, use three 400 µL aliquots to recover precipitates adhering to the sides of tubes and transfer them to a new 1.5 mL microcentrifuge tube. Note that this process is aided by using two P1000 pipettes and setting one to 425 µL and the other to 1 mL. Setting the pipette to 425 µL removes the need to pre-wet the pipette tip during the procedure. Briefly, following the removal of the final methanol wash, add 400 µL of RT 95% MeOH v/v to one of the sample tubes for each sample group and partially close the lids. Then, for one sample group at a time, use the second pipette to aspirate 200–400 µL of the methanol wash using the second pipette and carefully scrape down the sides of the tube while ejecting to dislodge precipitates. Typically, this step requires two complete passes around the inside of the tube. If pooling precipitates from multiple sample tubes, aspirate the entire volume, partially close the cap, and add it to the next tube to be pooled. After completing this process for all sample tubes of a given sample group, transfer the methanol wash and precipitates to a 1.5 mL microcentrifuge tube. For the remaining two aliquots of RT 95% MeOH v/v, closing the tube and mixing by vortex (3 s, maximum setting) is sufficient for collecting any remaining precipitates.
Non-crosslinked RNP fractions lack quantifiable amounts of protein and should not be used as a background control for UV-crosslinked RNP fractions due to the observation that recovery of non-crosslinked proteins by LEAP-RBP is dependent on RNA-bound proteins [18]. Nonetheless, for instances where they are included for comparison to UV-crosslinked samples, we recommend using a similar amount of RNA. For example, if UV-crosslinked samples containing > 0.25 µg of protein contain > 2.86 µg of RNA, use > 2.86 µg of RNA for non-crosslinked samples. Alternatively, use an equivalent percent fraction given a comparable amount of starting material.
For experiments where precision is key, we suggest increasing the number of aliquots used when preparing sample fractions and pooling prior to resuspension and/or after RNA and protein quantitation. For example, an RNP fraction prepared from one 200 µL aliquot of an AGP input suspension is less representative of the entire starting sample than an RNP fraction prepared from three 200 µL aliquots. Furthermore, processing each of the three 200 µL aliquots separately would help to reduce the variability introduced during specific steps of the LEAP-RBP procedure. This could include, for example, performing inversions/reversions for each aliquot independently by placing them on three separate sample racks. These types of measures do not replace the need for biologically independent replicates when studying biology-related phenomena, but they do help to reduce the likelihood that technical sources of variation (i.e., batch effects) impact biologically relevant observations.
Troubleshooting
Problem or issue Likely cause(s) Possible solution(s)
Tugging of the AGPC interphase when removing the aqueous phase during the repeated AGPC extraction procedure. Insufficient shearing of cell lysates during cell harvest. Centrifuge sample tubes prior to shearing, increase the number of passages when shearing lysates, increase the speed of passaging.
RNP pellets do not resuspend during the Turbo DNase digestion step. Attempting to add Turbo DNase Buffer to dry pellets. Complete resuspension of RNP pellets in TE buffer prior to adding Turbo DNase Buffer.
RNP pellets are brittle and break apart during 95% MeOH v/v washes. Centrifugation at 4 °C following inversions/reversions. Centrifuge samples at 20 °C.
Adding an excessive amount of Turbo DNase buffer during the DNA depletion step. Do not pre-wet pipette tips before adding pre-mixed master mix containing Turbo DNase buffer, Turbo DNase, and TE buffer to TE suspensions.
Forceful inversions during the washing procedure. Reduce speed and force when performing inversions.
Insufficient forceful reversion(s) prior to the centrifugation during the LEAP-RBP procedure. Using a sample tube rack, invert sample tubes until mixtures become displaced from the bottom of tubes and then perform a forceful reversion. Repeat the process in the opposite direction.
High free-protein recovery in LEAP-RBP fractions.
Incubating for longer than recommended after adding/layering precipitation solution to AGPC mixtures. Decrease the time to complete inversions/reversions after adding/layering the precipitation solution.
Not removing 95% MeOH v/v washes quickly after inverting tubes. Begin to remove the 95% MeOH v/v wash before the pellet is allowed to settle.
Attempting to invert and quickly remove 95% MeOH v/v washes from multiple sample tubes at the same time. Invert and quickly remove the 95% MeOH v/v wash from one tube at a time.
Insufficient denaturing of non-crosslinked RNA–protein and protein–protein interactors due to insufficient mixing of AGPC mixtures.
Mix samples after adding chloroform by vortex for at least 10 s at the highest speed setting.
Input samples and/or RNP pellets are not resuspended in 30 min using the recommended procedure. Attempting to dry a large number of precipitates or RNP pellets in the same sample tube. Spread out precipitates or RNP pellets so they dry more uniformly.
Colored flakes in samples. Degradation of graduation marks on serological pipettes after contacting phenol. Work quickly when transferring phenol using serological pipettes, < 2 min.
Plastic in samples. Melting of serological pipettes made of polystyrene after contacting chloroform or degradation of polypropylene tubes used to store chloroform for >1 h. Use pipette tips to transfer chloroform and decrease the time that chloroform is stored in polypropylene tubes to less than 1 h.
Unexpected decrease in LEAP-RBP yield. Chloroform stock has degraded. Replace the chloroform stock solution or increase the volume of chloroform and precipitation solution used to 15 µL and 860 µL, respectively.
RNP pellets extend up along the side of the tube after the first LEAP step. Not performing a sufficient number of inversions/reversions and/or not performing the final reversion with force. Increase the number of inversions/reversions to ten or more and increase the force and speed of the final reversion.
AGP input or final AGPC interphase suspensions contain more than 300 ng of RNA and DNA/µL. Dilute AGP input or final AGPC interphase suspensions with AGP.
RNP pellets appear scattered after the second LEAP step. Not maintaining TE-suspended RNPs at the bottom of tubes. Follow the recommended pipetting technique.
Insufficient mixing before and after adding chloroform. Ensure that Turbo DNase digests are mixed by vortex after adding 9 parts AGP (10 s, setting 5) and again after adding chloroform (10 s, maximum setting).
Inaccurate or widely variable RNA and protein quantitation data. Reagents have not been clarified, allowing particulates from the manufacturing processes to contaminate samples and react with the BCA reagent.
Ensure that reagents have been clarified and/or that reagents being used are of sufficient quality.
Large sample volumes have made the recommended clarification procedure inadequate. Aliquot the sample prior to clarification.
Trying to process too many samples in parallel during the clarification step and/or not transferring the clarified portion quickly after centrifugation. Reduce the number of sample suspensions being clarified and transfer the clarified portion quickly (< 5 min) after centrifugation.
Samples suspended in 1% LiDS TE have been stored at 4 °C or RT for extended periods of time before RNA and protein quantitation. Quantitate or store samples at -80 °C within 6 h of them being resuspended in 1% LiDS TE.
Phenol and/or GT have not been efficiently depleted from sample fractions during 95% MeOH v/v washes; for representative UV-spectrophotometric profiles of uncontaminated sample fractions, see Table S2. Remove additional supernatant following centrifugation during 95% MeOH v/v washes and/or increase the volume or number of 95% MeOH v/v washes used.
Acknowledgments
We thank members of the Nicchitta lab and in particular Rose Homoelle for helpful comments and critical feedback. Funding was provided by a grant from the NIH to CVN (GM139480).
Competing interests
The authors certify that they have no affiliations with or involvement in any organization or entity with any financial or non-financial interest in the subject matter and/or materials presented in this manuscript.
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Vieira-Vieira, C. H. and Selbach, M. (2021). Opportunities and Challenges in Global Quantification of RNA-Protein Interaction via UV Cross-Linking. Front Mol Biosci. 8: e669939.
Wheeler, E. C., Van Nostrand, E. L. and Yeo, G. W. (2017). Advances and challenges in the detection of transcriptome‐wide protein–RNA interactions. Wiley Interdiscip Rev RNA. 9(1): e1436.
Nechay, M. and Kleiner, R. E. (2020). High-throughput approaches to profile RNA-protein interactions. Curr Opin Chem Biol. 54: 37–44.
Van Nostrand, E. L., Freese, P., Pratt, G. A., Wang, X., Wei, X., Xiao, R., Blue, S. M., Chen, J. Y., Cody, N. A. L., Dominguez, D., et al. (2020). A large-scale binding and functional map of human RNA-binding proteins. Nature. 583(7818): 711–719.
Van Ende, R., Balzarini, S. and Geuten, K. (2020). Single and Combined Methods to Specifically or Bulk-Purify RNA–Protein Complexes. Biomolecules. 10(8): 1160.
Esteban‐Serna, S., McCaughan, H. and Granneman, S. (2023). Advantages and limitations of UV cross‐linking analysis of protein–RNA interactomes in microbes. Mol Microbiol. 120(4): 477–489.
Trendel, J., Schwarzl, T., Horos, R., Prakash, A., Bateman, A., Hentze, M. W. and Krijgsveld, J. (2019). The Human RNA-Binding Proteome and Its Dynamics during Translational Arrest. Cell. 176: 391–403.e19.
Queiroz, R. M. L., Smith, T., Villanueva, E., Marti-Solano, M., Monti, M., Pizzinga, M., Mirea, D. M., Ramakrishna, M., Harvey, R. F., Dezi, V., et al. (2019). Comprehensive identification of RNA–protein interactions in any organism using orthogonal organic phase separation (OOPS). Nat Biotechnol. 37(2): 169–178.
Urdaneta, E. C., Vieira-Vieira, C. H., Hick, T., Wessels, H. H., Figini, D., Moschall, R., Medenbach, J., Ohler, U., Granneman, S., Selbach, M., et al. (2019). Purification of cross-linked RNA-protein complexes by phenol-toluol extraction. Nat Commun. 10(1): 990.
Perez-Perri, J. I., Rogell, B., Schwarzl, T., Stein, F., Zhou, Y., Rettel, M., Brosig, A. and Hentze, M. W. (2018). Discovery of RNA-binding proteins and characterization of their dynamic responses by enhanced RNA interactome capture. Nat Commun. 9(1): 4408.
Castello, A., Fischer, B., Eichelbaum, K., Horos, R., Beckmann, B. M., Strein, C., Davey, N. E., Humphreys, D. T., Preiss, T., Steinmetz, L. M., et al. (2012). Insights into RNA Biology from an Atlas of Mammalian mRNA-Binding Proteins. Cell. 149(6): 1393–1406.
Baltz, A. G., Munschauer, M., Schwanhäusser, B., Vasile, A., Murakawa, Y., Schueler, M., Youngs, N., Penfold-Brown, D., Drew, K., Milek, M., et al. (2012). The mRNA-Bound Proteome and Its Global Occupancy Profile on Protein-Coding Transcripts. Mol Cell. 46(5): 674–690.
Kristofich, J. and Nicchitta, C. V. (2023). Signal-noise metrics for RNA binding protein identification reveal broad spectrum protein-RNA interaction frequencies and dynamics. Nat Commun. 14(1): 5868.
Barlow, J., Mathias, A., Williamson, R. and Gammack, D. (1963). A simple method for the quantitative isolation of undegraded high molecular weight ribonucleic acid. Biochem Biophys Res Commun. 13(1): 61–66.
Kristofich, J. and Nicchitta, C. (2024). High-throughput quantitation of protein-RNA UV-crosslinking efficiencies as a predictive tool for high confidence identification of RNA binding proteins. RNA. 30(6): 644–661.
Supplementary information
The following supporting information can be downloaded here:
Figure S1. Graphical representation of pipette tip wetting procedure.
Table S1. Compositional analysis of starting AGP input suspensions and dilutions calculator for downstream isolation of sample fractions.
Table S2. Dilution calculator for dilution of sample fractions prior to the clarification step.
Table S3. Compositional analysis of clarified sample fractions.
Video S1. Example of the LEAP-RBP inversion step being performed on eight 200 µL aliquots of an AGP suspension containing UV-crosslinked(0.4 J/cm2, 254 nm) HCT116 cells. Notably, samples were incubated for at least 1 min after the first nine inversions and 30 s after the final inversion.
Video S2. Example of the pre-wetting procedures used for aliquoting AGP input suspensions and chloroform precipitation solution.
Article Information
Publication history
Received: Mar 30, 2024
Accepted: Jun 19, 2024
Available online: Jul 5, 2024
Published: Jul 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Biochemistry > RNA > RNA-protein interaction
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53 | https://bio-protocol.org/en/bpdetail?id=53&type=1 | # Bio-Protocol Content
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This is an In Press version of the protocol that has not yet been assigned to an issue.
Peer-reviewed
Standard PCR Protocol
Fanglian He
In Press
Published: Apr 5, 2011
DOI: 10.21769/BioProtoc.53 Views: 39916
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Abstract
This protocol describes basic steps of a PCR experiment using home-made Taq DNA polymerase. Some steps may vary with different DNA polymerase.
Materials and Reagents
Tris-HCl (Sigma-Aldrich)
KCl (EM SCIENCE)
MgCl2 (EM SCIENCE)
Gelatin (Sigma-Aldrich)
Taq DNA polymerase (home-made)
dNTPs (New England Biolabs, catalog number: N0447L )
Template DNA (genomic, plasmid, cosmid, bacterial/yeast colony, etc.)
Primers
Equipment
Thermal cycler (MJ Research)
Procedure
Prepare DNA template:
Usually, for plasmid DNA, 1-10 ng; for genomic DNA, 50-100 ng per reaction is needed. Normally, DNA template does not need to be purified. However, both purity and the amount of template can strongly influence the outcome of the reaction.
Design primer:
Generally, primers used are 18-23 mer in length. Use Primer3 free online software (reference 1) to design primers.
Determine annealing temperature:
Melting temperature (Tm) of primers can be calculated by the following formula: Tm = [(#of A + T residues) x 2] + [(#of G + C residues) x 4] °C. Tm-5 °C is a good annealing temperature to start with. However, optimal annealing temperatures can only be determined experimentally for a certain primer/template combination. Temperature gradient PCR is often a way to finalize an optimal annealing temperature.
Prepare 10x PCR reaction buffer, include:
100 mM Tris-HCl (pH 8.3)
500 mM KCl
15 mM MgCl2
0.1% gelatin
Note: The MgCl2 concentration is typically 10-15 mM. However, the optimum concentration needs to be determined experimentally. Mg2+ forms a soluble complex with dNTP's which facilitates dNTP incorporation, and stimulates polymerase activity. It also promotes and stabilizes primer and template interaction. Thus, Increasing the magnesium concentration has the same effect as lowering the annealing temperature. Too low Mg2+ leads to low yields (or no yield) and too much Mg2+ cause nonspecific products.
For a 100 μl reaction, add:
10x PCR buffer 10 μl
DNA template (5 ng μl-1) 1 μl
Primer A (50 mM) 1 μl
Primer B (50 mM) 1 μl
dNTPs (2 mM) 10 μl
Taq (5 U μl-1) 1 μl
Sterile ddH2O 76 μl
Notes:
For some PCR machines that do not have a heated lid, mineral oil needs to be added to each reaction to prevent evaporation of the sample.
Prepare a control reaction with no template DNA and an additional 10 μl of sterile water.
A typical PCR program may be:
Initial denaturation, 4-8 min at 94-95 °C.
Denaturation, 15 sec at 94-95 °C.
Annealing, 15 sec at x °C (depends on Tm).
Extension, x sec (depends on product length, 1 min kb-1) at 72 °C.
Return to step 2 for 30-35 additional cycles.
Final extension, 10 min at 72 °C.
Keep sample at 4 °C until loading.
References
http://frodo.wi.mit.edu/primer3/
Article Information
Copyright
© 2011 The Authors; exclusive licensee Bio-protocol LLC.
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Category
Molecular Biology > DNA > PCR
Molecular Biology > DNA > DNA cloning
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531 | https://bio-protocol.org/en/bpdetail?id=531&type=0 | # Bio-Protocol Content
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Peer-reviewed
Fear Conditioning Assay in Mouse
MW Melissa Wang
IM Isabel A. Muzzio
Published: Vol 3, Iss 7, Apr 5, 2013
DOI: 10.21769/BioProtoc.531 Views: 13852
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Original Research Article:
The authors used this protocol in The Journal of Neuroscience Nov 2012
Abstract
The study of fear memory is important for understanding various anxiety disorders in which patients experience persistent recollections of traumatic events. These memories often involve associations of contextual cues with aversive events; consequently, Pavlovian classical conditioning is commonly used to study contextual fear learning. A form of contextual fear conditioning that is becoming increasingly important as an animal model of anxiety disorders uses predator odor as a fearful stimulus. Innate fear responses to predator odors are well characterized and reliable; however, attempts to use these odors as unconditioned stimuli in fear conditioning paradigms have been highly dependent on experimental setup and have produced inconsistent behavioral results. Here we present a contextual fear conditioning paradigm using coyote urine as the unconditioned stimulus, which has been shown to produce consistent contextual freezing in response to fear learning (Wang et al., 2012).
Materials and Reagents
Mouse C57BL/6, male, 6 weeks to 4 months old (we suggest using mice of a similar age, with ~3 months old being optimal) (Jackson Laboratories)
Paper towel, cut into 2.5 by 2.5 cm squares
Scotch tape
100% coyote urine (Maine Outdoor Solutions, catalog number: ACOYD )
Ethanol
Cleaning solution (409 and/or Clorox Clean-Up)
Liquid soap
Equipment
Cylindrical Plexiglas training context 35 cm in diameter and 35 cm tall with a platform base and fitted removable cylindrical wall (custom made by Just Plastics), painted white with 5-6 black distinct visual cues along the wall in basic shapes, each about 10-12 cm in width.
Camera/computer setup to record animal movement – overhead camera with low lighting levels: Bright enough that the computerized tracking system can track the mouse, but dark enough that the mouse is comfortable exploring the environment.
Limelight (Actimetrics) or some other such behavioral tracking system
Procedure
Handle mouse for 5 min per day for 3 consecutive days prior to beginning the experiment. Handle by holding each mouse and letting it climb around on and explore your hands so that they habituate to the experimenter. Mice can be run successively in groups of 5, but housed individually.
One day before fear conditioning, place the mouse in the training context for 10 min to allow habituation to the context. Fold a piece of tape into a loop and affix to the underside of a paper towel square, taping it to the center of the cylindrical environment. Saturate the paper towel with 20 drops of water before placing the mouse into the context. For contextual consistency, present a paper towel square wetted with water in all sessions except for the predator odor conditioning session. Record all sessions on a computer with Limelight or the behavioral tracking system of your choice.
The next day, expose the mouse to the context for 10 min to take a baseline freezing measure (again placing water in the center of the context). Remove the mouse from the context, and replace the water with a paper towel square saturated with 20 drops of 100% coyote urine. Place the mouse back in the context in the presence of coyote urine for 5 min. Remove the mouse from the contex (as a control, expose other mice to just water in place of the predator odor).
Clean the base of the context thoroughly with 409 and/or Clorox Clean-up and soap several times. Dry and wipe with ethanol (also use ethanol to clean the context between all sessions). Ensure that no coyote odor remains. Meanwhile, air out the conditioning room by opening the door and turning on several fans. If only water has been used, clean the context in the same manner to ensure consistency.
One hour after the conditioning session, give a short-term memory retention test by placing the mouse again in the cleaned context with water for 10 min.
Perform 10 min long-term memory retention tests at 24 h intervals after the conditioning session. This can be repeated for as many days as you wish to continue the study.
Analyze the percentage of time spent freezing for each session. In Limelight, you can export velocity measures for each second and use Excel filters to calculate what percentage of the total time the mouse spent below a certain freezing threshold velocity (0.5 cm/sec or something to that effect).
Acknowledgments
This protocol is adapted from Wang et al. (2012).
References
Wang, M. E., Wann, E. G., Yuan, R. K., Ramos Alvarez, M. M., Stead, S. M. and Muzzio, I. A. (2012). Long-term stabilization of place cell remapping produced by a fearful experience. J Neurosci 32(45): 15802-15814.
Article Information
Copyright
© 2013 The Authors; exclusive licensee Bio-protocol LLC.
How to cite
Readers should cite both the Bio-protocol article and the original research article where this protocol was used:
Wang, M. E. and Muzzio, I. A. (2013). Fear Conditioning Assay in Mouse. Bio-protocol 3(7): e531. DOI: 10.21769/BioProtoc.531.
Wang, M. E., Wann, E. G., Yuan, R. K., Ramos Alvarez, M. M., Stead, S. M. and Muzzio, I. A. (2012). Long-term stabilization of place cell remapping produced by a fearful experience. J Neurosci 32(45): 15802-15814.
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Category
Neuroscience > Behavioral neuroscience > Learning and memory
Neuroscience > Behavioral neuroscience > Animal model > Mouse
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55 | https://bio-protocol.org/en/bpdetail?id=55&type=1 | # Bio-Protocol Content
Improve Research Reproducibility
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This is an In Press version of the protocol that has not yet been assigned to an issue.
Peer-reviewed
Common Worm Media and Buffers
Fanglian He
In Press
Published: Apr 5, 2011
DOI: 10.21769/BioProtoc.55 Views: 53835
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Abstract
Here are recipes of some media and solutions often used in C. elegans research.
Materials and Reagents
Agar, peptone (BD Biosciences)
Cholesterol (Sigma-Aldrich)
Streptomycin (Sigma-Aldrich)
Nystatin (Life Technologies, Gibco®)
Bleach (Clorox)
Potassium phosphate
Clorox bleach
NaCl
CaCl2
MgSO4
EtOH
FeSO4.7H2O
Na2EDTA
MnCl2.4H2O
ZnSO4.7H2O
CuSO4.5H2O
KH2PO4
Na2HPO4
Equipment
60 x 15 mm plate
Plastic boxes
Recipes
Nematode growth medium (NGM) agar: For the maintenance of worms.
For 1 liter medium
3 g NaCl
17 g agar
2.5 g peptone
1 ml cholesterol (5 mg ml-1 in 95% EtOH)
975 ml H2O
Autoclave, and then add the following sterile solution (autoclaved)
1 ml 1 M CaCl2
1 ml 1 M MgSO4
25 ml 1 M potassium phosphate (pH 6) (to avoid precipitation, mix between addition of MgSO4 and potassium phosphate)
To make 1 M potassium phosphate (pH 6): For 1 liter, dissolve 136.1 g KH2PO4 in about 800 ml dH2O, then adjust to pH 6.0 with solid KOH (approx 15 g) before bringing up to volume. Make 100 ml aliquots and autoclave.
Need to add streptomycin (300 ng ml-1) if plate is used for seeding bacterial food E coli OP50-1. Typically pour 60 x 15 mm plate and store NGM plates in plastic boxes with covers at room temperature.
S-basal medium (adapted from the Kim Lab at Stanford) : For liquid culture of worms.
For 1 liter medium
5.8 g NaCl
50 ml 25 ml 1 M potassium phosphate (pH 6)
1 ml cholesterol (5 mg ml-1 in 95% EtOH)
950 ml dH2O
Autoclave, and then add the following sterile solution (autoclaved)
3 ml 1 M CaCl2
3 ml 1 M MgSO4
10 ml trace metals solution
10 ml 1 M potassium citrate (pH 6.0)
10 ml 100x Nystatin (antifungal agent, keep in freezer; do not have to add it all the time).
To make 500 ml trace metals solution
0.346 g FeSO4.7H2O
0.930 g Na2EDTA
0.098 g MnCl2.4H2O
0.144 g ZnSO4.7H2O
0.012 g CuSO4.5H2O
Sterilize by autoclaving. Keep in dark (wrap in foil).
To make 100 ml of 1 M potassium citrate: dissolve 21.02 g citric acid, monohydrate in 80 ml and adjust to pH 6.0 with solid KOH (approx 17g) before bringing up to volume.
Worm M9 buffer
3 g KH2PO4
6 g Na2HPO4
5 g NaCl
Add H2O to 1 liter. Sterilize by autoclaving.
After solution cools down, add 1 ml autoclaved/sterile 1 M MgSO4.
100 ml 2x worm lysis solution: For worm egg prep
50 ml ddH2O
10 ml 10 M NaOH
40 ml Clorox bleach
Make fresh and store at 4 °C up to one week.
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© 2011 The Authors; exclusive licensee Bio-protocol LLC.
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56 | https://bio-protocol.org/en/bpdetail?id=56&type=1 | # Bio-Protocol Content
Improve Research Reproducibility
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This is an In Press version of the protocol that has not yet been assigned to an issue.
Peer-reviewed
Synchronization of Worm
Fanglian He
In Press
Published: Apr 5, 2011
DOI: 10.21769/BioProtoc.56 Views: 24566
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Abstract
Attaining a large synchronized population of worms is desirable for use in some assays in order to eliminate variation in results due to age differences. Two ways to get synchronized worms include egg preparation via bleaching and egg lay. The former in general yields more progeny than the latter, however, egg lay can generate a better synchronized population than egg preparation.
Materials and Reagents
General chemicals (Sigma-Aldrich)
Regular bleach (Clorox)
M9 buffer
Worm lysis solution
Equipment
Votex (Scientific Industries)
Vacuum pump (Welch)
15 ml conical tube
60 x 15 mm petri dish (VWR)
Procedure
Egg preparation via bleaching
Chunk starved worms onto a seeded (with bacterial food, E coli OP50-1) NGM plate (60 x 15 mm). Allow the worms to grow for about 2 days at 25 °C.
Once you have plenty of eggs/adults, pour 6 ml of M9 buffer onto the plate and gently swirl it to dislodge the worms.
Transfer 5 ml worms to a 15 ml conical tube.
Add 5 ml of 2x worm lysis solution to the tube.
Vortex the tube at max speed for approximately 4 min or until you see very few intact adult worms (usually bleach no longer than 6 min, otherwise the eggs will be killed).
Centrifuge at ~1,000 x g for 1 min.
Carefully decant the supernatant without disturbing the worm pellet.
Wash twice with 10 ml of M9 buffer (spin at ~1,000 x g for 1 min)
Add 7 ml of M9 buffer to resuspend the egg pellet.
Let eggs shake overnight at 20 °C to hatch. Since there is no food the larvae should be halted at the L1 stage.
Next day, let the tube sit on the bench for 10 - 15 min and starved L1 worms can precipitate by gravity.
Remove most of supernatant and distribute the liquid onto seeded plates or into liquid culture.
Note: This protocol is also used to remove bacterial and yeast contamination from a worm strain.
Egg lay
Gravid worms have a maximal egg-laying rate of 7 eggs/h at 20 °C.
Pick 10-15 gravid worms/plate.
Incubate at desired temperature (15, 20, or 25 °C) for desired time (2-6 h).
Remove adults by suction off the plate or picking.
Grow until progeny are at appropriate stage.
References
Sulston, J. & Hodgkin, J. (1988) Methods. In: The Nematode Caenorhabditis elegans. (Ed.): W.B. Wood. Cold Spring Harbor Laboratory Press: New York, 587-606.
Shapira, M. and Tan, M. W. (2008). Genetic analysis of Caenorhabditis elegans innate immunity. Methods Mol Biol 415: 429-442.
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569 | https://bio-protocol.org/en/bpdetail?id=569&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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Peer-reviewed
Stem Microsome Preparation and Mannan Synthase Activity Assay
Yan Wang
Published: Vol 3, Iss 10, May 20, 2013
DOI: 10.21769/BioProtoc.569 Views: 8385
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Original Research Article:
The authors used this protocol in The Plant Journal Jan 2013
Abstract
Mannans are hemicellulosic polysaccharides and are present in cell walls of all land plants. Mannan polysaccharides are synthesized by two enzymes, mannan synthase (ManS) for backbone (mannan or glucomannan) synthesis and galactomannan galactosyl transferase for side-chain (galactosyl) addition. Here, a method for ManS activity assay using microsomes freshly isolated from Arabidopsis stems is described. This method can be applied to isolation of microsomes from any tissues of Arabidopsis or any other plants.
Keywords: Arabidopsis Cell wall Microsomes Galactomannan Mannan synthase
Materials and Reagents
cOmplete, Mini, EDTA-free Protease Inhibitor Cocktail Tablets (Roche, catalog number: 11836170001 )
GDP-[14C]-Man (9.694 GBq/mmol, 262.0 mCi/mmol, 76.34 μM) (PerkinElmer, catalog number: NEC536050UC )
Bicinchoninic acid (BCA) Protein Assay Kit (Pierce Antibodies, catalog number: 23225 or 23227 )
OptiPhase Supermix Cocktail (PerkinElmer, catalog number: 1200-439 )
GDP-Man (Sigma-Aldrich, catalog number: G-7377 )
Carob galactomannan (Megazyme International, catalogue number: P-GALML )
Sucrose
MgCl2
Extraction buffer (EB) (see Recipes)
4x ManS assay buffer (see Recipes)
1 mM GDP-Man (see Recipes)
1% carob galactomannan (see Recipes)
Equipment
Mortar and pestle
13 ml Sarstedt tube (Sarstedt, catalog number: 60.540.500 )
Centrifuge
4 ml plastic liquid scintillation counting vial (PerkinElmer, catalog number: 1200-421 )
1450 MicroBeta TriLux Microplate Scintillation and Luminescence Counter (PerkinElmer, catalog number: 1450-024 )
Procedure
Grow Arabidopsis plants in a growth chamber under standard growth conditions (20 °C, 100 μmol/m2/s, 16 h light/8 h dark, 60% humidity) for 6-7 weeks.
Harvest the whole stem (including secondary stems) from a single plant, and remove leaves, flowers and siliques. The stem should be big enough (at least 200 mg of fresh weight) but not very old or hard (without any yellow siliques).
Weigh the stem and cut it into approximately 1 cm segments.
Immediately transfer the stem segments into a mortar pre-chilled on ice, and add ice-cold EB (~1 ml EB/100 mg stem).
Grind the stem segments in EB on ice with a mortar and pestle until the tissue is homogenized well (~5 min).
I usually grind the tissue in the cold room.
Transfer stem homogenate to a pre-chilled 13 ml Sarstedt tube using a 1 ml wide-bore pipetman tip (with the sharp end cut off), and centrifuge it at 3,000 x g at 4 °C for 10 min.
Transfer the supernatant to a new pre-chilled 13 ml Sarstedt tube, and centrifuge it at 17,000 x g at 4 °C for 20 min.
Centrifuge the supernatant from step 7 at 100,000 x g at 4 °C for 90 min to pellet microsome membranes.
Resuspend the membrane pellet in EB (0.5 μl EB/mg stem) by pipetting up and down using a 200 μl tip. The membranes are pelleted tightly, so it takes some time to resuspend the pellet. I usually do it in the cold room.
Aliquot a small volume of the microsome membranes, diluted it by 10 fold with EB, and store it at –20 °C. The diluted sample will be used for quantifying protein concentration using the BCA Protein Assay Kit.
Transfer microsome suspension to a 1.5 ml Eppendorf tube, vortex briefly, and incubate on ice for 5 min to let large particulates settle down.
Aliquot 20 μl of microsomes (do not pipet the precipitate on the bottom) into 1.5 ml Eppendorf tubes pre-chilled on ice. Boil 2-3 tubes of microsomes at 100 °C for 10 min, and the boiled samples will be used as a boiled control.
Prepare reaction cocktail by adding and mixing the following reagents.
4x ManS assay buffer
10 μl
(final concentration in reaction: 1x)
1 mM GDP-Man
0.85 μl
(final concentration in reaction: 21.18 μM)
GDP-[14C]-Man(76.34 μM)
2 μl
(final concentration in reaction: 3.82 μM)
Deionized H2O
7.15 μl
Make master mix by scaling up the volumes based on the numbers of reaction tubes.
Pipet 20 μl of the reaction cocktail into each tube containing 20 μl of microsomes, and briefly vortex to mix. Allow ~5 sec staggering time between two samples.
Conduct reactions at room temperature for 1 h.
Stop the reactions by adding 1 ml of 70% ethanol containing 2 mM EDTA and 10 μl of 1% carob galactomannan. Precipitate assay products at –20 °C for at least 1 h.
Pellet the assay products by centrifuging at 16,000 x g at 4 °C for 10 min.
Wash the pellets with 70% ethanol containing 2 mM EDTA four times, and repeat centrifugation after each wash.
Resuspend each washed pellet in 300 μl of water.
Transfer suspension to a 4 ml plastic scintillation vial, and then add 3 ml OptiPhase Supermix Cocktail to the vial. Cap the vial and mix the sample well by vortexing briefly.
Perform liquid scintillation counting using a 1450 MicroBeta TriLux Microplate Scintillation and Luminescence Counter.
Calculate the in vitro ManS enzymatic activities (shown as picomoles of GDP-Man incorporation per hour per mg protein) based on radioactivity of the assay products using the following formula.
Specific ManS activity = (CPM ÷ counting efficiency) ÷ 2,220,000 ÷ (GDP-[14C]-Man radioactivity shown as μCi/μmol) x 1,000,000 x [(cold GDP-Man concentration + hot GDP-Man concentration) ÷ hot GDP-Man concentration] ÷ 1 h ÷ [20 x (protein concentration as μg/μl) ÷ 1,000]
Detailed explanations are as follows:
DPM (decaying per minute) = CPM (counting per minute) ÷ counting efficiency
(This formula is used to convert the detected radioactivity (cpm) of the assay products to the actual radioactivity (dpm) based on the detection efficiency of the of the liquid scintillation counting machine)
Hot GDP-Man incorporation (pmol) = DPM ÷ 2,220,000 dpm/μCi ÷ (GDP-[14C]-Man radioactivity shown as μCi/μmol) x 1,000,000 pmol/μmol
(This formula is used to convert the radioactivity (dpm) of the assay products to pmol radio-labeled (hot) GDP-Man incorporation. Note the unit conversions: 1 μCi = 2,220,000 dpm; 1 μmol = 1,000,000 pmol.)
Total GDP-Man incorporation (pmol) = hot GDP-Man incorporation (pmol) x [(cold GDP-Man concentration + hot GDP-Man concentration) ÷ hot GDP-Man concentration]
[The total GDP-Man incorporation (pmol) is calculated by dividing hot GDP-Man incorporation (pmol) by the ratio of the hot GDP-Man concentration to the total (hot + cold) GDP-Man concentration]
Protein mass (mg) of 20 μl microsomes = 20 μl x (protein concentration as μg/μl) ÷ 1,000 μg/mg
Specific ManS activity (pmol GDP-Man incorporation/h/mg protein) = total GDP-Man incorporation (pmol) ÷ 1 h ÷ protein mass (mg)
For cold and hot GDP-Man reagents used in this protocol:
Specific ManS activity = (CPM ÷ counting efficiency) ÷ 2,220,000 ÷ 262 x 1,000,000 x [(21.18 + 3.82) ÷ 3.82] ÷ 1 h ÷ [20 x (protein concentration as μg/μl) ÷ 1,000]
Recipes
Extraction Buffer (EB)
50 mM HEPES-KOH, pH 7.5
0.4 M Sucrose
10 mM MgCl2
Filter sterilize and store at 4 °C
Before use, add 1 cOmplete, Mini, EDTA-free Protease Inhibitor Cocktail Tablet per 10 ml buffer.
4x ManS Assay Buffer
200 mM HEPES-KOH (pH 7.5)
10 mM DTT
10 mM MgCl2
20 mM MnCl2
24% glycerol
Store at –20 °C
1 mM GDP-Man
Dissolve 5 mg GDP-Man in 770 μl of 10 mM HEPES-KOH (pH 7.5) to a final concentration of 10 mM. Dilute a small volume of 10 mM GDP-Man to 1 mM with H2O
1% carob galactomannan
Dissolve 100 mg carob galactomannan in H2O with a final volume of 10 ml. Heat to facilitate dissolution of galactomannan
Acknowledgments
This protocol is adapted from Wang et al. (2013).
References
Wang, Y., Mortimer, J. C., Davis, J., Dupree, P. and Keegstra, K. (2013). Identification of an additional protein involved in mannan biosynthesis. Plant J 73(1): 105–117.
Article Information
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How to cite
Wang, Y. (2013). Stem Microsome Preparation and Mannan Synthase Activity Assay. Bio-protocol 3(10): e569. DOI: 10.21769/BioProtoc.569.
Download Citation in RIS Format
Category
Biochemistry > Carbohydrate > Polysaccharide
Biochemistry > Protein > Activity
Cell Biology > Organelle isolation > Microsome
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57 | https://bio-protocol.org/en/bpdetail?id=57&type=1 | # Bio-Protocol Content
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Lifespan Assay
Fanglian He
In Press
Published: Apr 5, 2011
DOI: 10.21769/BioProtoc.57 Views: 27450
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Abstract
This assay is used to address aging-related questions in worms.
Materials and Reagents
NGM medium
5-fluorodeoxyuridine (FUDR) (Sigma-Aldrich, catalog number: F0503 )
Equipment
Incubators
NGM agar plates (35 x 10 mm)
60 x 15 and 35 x 10 mm petri dish (VWR)
Procedure
Synchronize worms by either egg prep or egg lay.
Animals are grown on NGM plates until they reach the L4 stage (about 36 h at 25 °C).
Make small NGM agar plates (35 x 10 mm) containing 5-fluorodeoxyuridine (FUDR; 0.1 mg ml-1) to prevent growth of progeny. Make plates fresh and store in the dark at 4 °C up to one week.
Seed a FUDR-containing plate fresh. Spot overnight OP50-1 culture (50 μl of 20x concentrated culture) onto FUDR plates one day before transferring worms. Keep seeded plates at RT overnight.
Transfer 30 L4 to each FUDR plate and 4-5 plates for each strain.
Grow at desired temperature (15, 20, or 25 °C). Animals are scored every 1 to 3 days subsequently and scored as dead when they no longer respond to gentle prodding with a platinum wire. Remove the dead worms after counting. Worms found dehydrated on plate walls are not counted as dead worms.
Lifespan is defined as the day animals are at the L4 larval stage (time t = 0) until the day they are scored as dead.
References
Wolkow, C. A., Kimura, K. D., Lee, M. S. and Ruvkun, G. (2000). Regulation of C. elegans life-span by insulinlike signaling in the nervous system. Science 290(5489): 147-150.
Apfeld, J. and Kenyon, C. (1999). Regulation of lifespan by sensory perception in Caenorhabditis elegans. Nature 402(6763): 804-809.
Mitchell, D. H., Stiles, J. W., Santelli, J. and Sanadi, D. R. (1979). Synchronous growth and aging of Caenorhabditis elegans in the presence of fluorodeoxyuridine. J Gerontol 34(1): 28-36.
Article Information
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Category
Developmental Biology > Cell growth and fate
Cell Biology > Cell viability > Cell proliferation
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58 | https://bio-protocol.org/en/bpdetail?id=58&type=1 | # Bio-Protocol Content
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Making Males of C. elegans
Fanglian He
In Press
Published: Apr 20, 2011
DOI: 10.21769/BioProtoc.58 Views: 28250
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Abstract
Getting males from a hermaphrodite population. This is a modified version of protocol originally written by Michael Koelle at Yale University.
Materials and Reagents
NGM plates
Equipment
25 and 30 °C incubators
Procedure
Set up ~6 NGM plates (seeded with normal bacterial food E. coli OP50-1) with 10 L4 hermaphrodites each.
Heat shock 4-6 h (no more than 6 h) at 30 °C.
Move plates to 25 °C. You should get a few males per plate in the F1 generation.
Note: To get more males, it is best to set them up with an excess of L4 hermaphrodites to ensure more males recover in the next generation.
Male stocks should be maintained by selecting out an excess of males, along with a few L4 hermaphrodites in each generation (usually, male to hermaphrodite ratio is about 1: 10).
Article Information
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© 2011 The Authors; exclusive licensee Bio-protocol LLC.
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Category
Developmental Biology > Cell growth and fate
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59 | https://bio-protocol.org/en/bpdetail?id=59&type=1 | # Bio-Protocol Content
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RNA Interference (RNAi) by Bacterial Feeding
Fanglian He
In Press
Published: Apr 20, 2011
DOI: 10.21769/BioProtoc.59 Views: 27551
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Abstract
There are 3 ways to perform RNAi in worms: microinjection, soaking and feeding. In the feeding protocol, RNAi is induced by cultivating worms on bacteria expressing gene-specific dsRNA. dsRNA is expressed in E. coli and ingested by worms. This protocol describes the feeding protocol to induce RNAi.
Materials and Reagents
C. elegans RNAi clone/libraries (Open Biosystems or Source BioScience LifeSciences)
Ampicillin (IBI Scientific)
Tetracycline
IPTG (Gold Biotechnology)
LB agar medium: Agar (BD Biosciences), tryptone (BD Biosciences), yeast extract (BD Biosciences), NaCl (Research Organics)
RNAi plate (NGM/IPTG/Ampicillin) (see Recipes)
Equipment
Aluminum foil
Petri dish (35 x 10 mm)
Procedure
Streak dsRNA-expressing E coli onto LB agar plate containing ampicillin (50 μg ml-1) and tetracycline selection (12.5 μg ml-1) and incubate at 37 °C overnight.
Note: There are two RNAi feeding libraries. One was constructed by the Vidal and Heuvel lab and can be ordered from Open Biosystems; the other was constructed by Julie Ahringer's lab and is available at Source BioScience LifeSciences.
Inoculate bacteria in 3 ml LB liquid medium containing ampicillin (100 μg ml-1) only and incubate at 37 °C overnight.
Spin down all 3 ml culture and pour off supernatant to 150 μl (concentrate culture by 20x). Resuspend pellet.
Transfer 50 μl of cell resuspend to center of RNAi plate (NGM/IPTG/Ampicillin). Let dry (wrapped in aluminum foil) and induce overnight at room temperature (RNAi-seeded plates can be stored at RT for 2-3 days before use).
Place 10-15 egg-laying worms on each plate. Incubate 2 - 6 h at 20 or 25 °C. Suck off parents and incubate at desired temperature until desired stage for further experiments.
Note: Instead of egg lay, synchronize worms by bleach and transfer starved L1 larva to RNAi plates.
Recipes
RNAi plate (NGM/IPTG/Ampicillin)
Use the same recipe for making NGM agar medium but instead of adding streptomycin, add IPTG to final a concentration of 1 mM and ampicillin with a concentration of 100 μg ml-1. Typically pour onto small petri dish (35 x 10 mm).
References
Timmons, L. and Fire, A. (1998). Specific interference by ingested dsRNA. Nature 395(6705): 854.
Rual, J. F., Ceron, J., Koreth, J., Hao, T., Nicot, A. S., Hirozane-Kishikawa, T., Vandenhaute, J., Orkin, S. H., Hill, D. E., van den Heuvel, S. and Vidal, M. (2004). Toward improving Caenorhabditis elegans phenome mapping with an ORFeome-based RNAi library. Genome Res 14(10B): 2162-2168.
Kamath, R. S. and Ahringer, J. (2003). Genome-wide RNAi screening in Caenorhabditis elegans. Methods 30(4): 313-321.
Article Information
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© 2011 The Authors; exclusive licensee Bio-protocol LLC.
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Category
Molecular Biology > RNA > RNA interference
Molecular Biology > RNA > Transfection
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60 | https://bio-protocol.org/en/bpdetail?id=60&type=1 | # Bio-Protocol Content
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Single Worm PCR
Fanglian He
In Press
Published: Apr 20, 2011
DOI: 10.21769/BioProtoc.60 Views: 20826
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Abstract
This protocol is used for genotyping single worms.
Materials and Reagents
Nonidet P-40 (Sigma-Aldrich, catalog number: 74385 )
Gelatin (Sigma-Aldrich)
Proteinase K (Promega Corporation, catalog number: V3021 )
KCl
Tris (pH 8.3)
MgCl2
Nonidet P-40
Tween-20
Gelatinin
Lysis buffer (see Recipes)
Equipment
PCR tubes
Procedure
Transfer one adult worm directly from a plate to 5 μl of lysis buffer in each PCR tube.
Note: To avoid transferring a large amount of bacteria, pick worm from region away from bacterial lawn. Also, try not to pick an old worm or starving worm.
Spin capped PCR tubes briefly to bring down worm to the bottom of the tube.
Freeze at -80 °C for 10 min or longer (up to a week).
Lyse the worms for release of genomic DNA.
Heat sample at 60 °C for 1 h, then inactivate protease K at 95 °C for 15 min. Store the worm DNA at -80 °C (or -20 °C for temporary storage) if needed.
Perform standard PCR reaction, use 5 μl worm DNA as template for 50 μl reaction.
Recipes
Lysis buffer
50 mM KCl
10 mM Tris (pH 8.3)
2.5 mM MgCl2
0.45% Nonidet P-40
0.045% Tween-20
0.01% (w/v) gelatinin
Autoclave and store at 4 °C (good for more than 6 months) or -20 °C for long-term storage. Right before use, add proteinase K stock to the lysis buffer with final concentration 60 μg ml-1.
References
Williams, B. D., Schrank, B., Huynh, C., Shownkeen, R. and Waterston, R. H. (1992). A genetic mapping system in Caenorhabditis elegans based on polymorphic sequence-tagged sites. Genetics 131(3): 609-624.
Article Information
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© 2011 The Authors; exclusive licensee Bio-protocol LLC.
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Category
Molecular Biology > DNA > PCR
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61 | https://bio-protocol.org/en/bpdetail?id=61&type=1 | # Bio-Protocol Content
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Affinity Purification of Yeast Protein-interacting Metabolites for ESI-MS Analysis
Xiyan Li
In Press
Published: Apr 20, 2011
DOI: 10.21769/BioProtoc.61 Views: 15239
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Abstract
The method described here can be used to discover in vivo protein-metabolite interactions. Metabolite-protein complexes are purified from yeast cell lysates by an affinity tag that recognizes the protein of interest. The protein-bound metabolites are extracted for identification by mass spectrometry, while the protein is concurrently analyzed by gel electrophoresis. A parallel experiment using cell lysate without target protein should be used as a negative control. The metabolite extract should be analyzed within 1-2 days to avoid undesired chemical reaction.
Keywords: Protein purification Affinity purification Yeast Metabolite extraction Metabolomics
Materials and Reagents
Cells
1x phosphate buffered saline (PBS)
Methanol (mass spec grade)
Water (mass spec grade)
2x laemmli buffer (for SDS-page)
NH4Ac
EGTA
DTT
PMSF
Roche protease inhibitor tablets (Roche Diagnostics)
Lysis buffer (see Recipes)
Wash buffer 1 (see Recipes)
Wash buffer 2 (see Recipes)
Equipment
Zirconia silica beads (Bio Spec Products)
Rabbit IgG-conjugated dynabeads
Eppendorf protein Lobind tubes
FastPrep cell lyser with an adapter for 2 ml tubes
Hula mixer (Life Technologies, InvitrogenTM) or similar product
Magnetic stand for 1.5/2.0 ml tubes
Heat block
Procedure
Add equal volume of 0.5 mm Zirconia silica beads (stored at -20 °C) to cells from 150 ml culture, add in 950 μl lysis buffer, homogenate on FastPrep 24, 3 x 40 sec min at 6.5 m/sec with 2 min interval on ice.
Note: Wash the IgG Dynabeads 2x in 1x PBS, 3x in lysis buffer. Re-suspend in lysis buffer. Use 50 μl per sample.
Spin down lysate at 14,000 rpm, 10 min, and transfer supernatant (lysate) to 2.0 ml Lobind tubes. Store at 4 °C.
Add 950 μl lysis to the cell pellet and lyse again as step 1.
Repeat step 2 and combine the lysate. Add 50 μl IgG beads.
Incubate 30 min at 4 °C with end-over-end invertion on Hula mixer.
Use magnetic stand to separate beads from lysate.
Wash the beads in 0.8 ml wash buffer 1, and 0.8 ml in wash buffer 2. Transfer beads with wash buffer 2 to a new tube. Each time buffer is added to the beads, invert at 4 °C until homogenate, briefly spin down the beads (10 sec), put on magnetic stand for at least 30 sec, and pipet off the buffer.
Add 50 μl methanol (MS grade) to the beads, pipette mix, 15 min at room temperature, and separate on magnetic beads, repeat and combine the methanol extracts in MS vials.
Add 30 μl 2x SDS sample buffer to the beads, boil 15 min, load 15 μl on SDS-page for protein yield evaluation.
Recipes
Lysis buffer
200 mM NH4Ac (stock: 5 M)
1 mM EGTA (stock: 500 mM EGTA)
1 mM DTT (stock: 1 M)
1 mM PMSF (stock: 100 mM in ethanol, 4 °C)
Roche Protease inhibitor tablets (1x), if no EDTA, add to final 1 mM
Wash buffer 1
500 mM NH4Ac (stock: 5 M) ligation reaction
Wash buffer 2
50 mM NH4Ac (stock: 5 M)
References
Li, X., Gianoulis, T. A., Yip, K. Y., Gerstein, M. and Snyder, M. (2010). Extensive in vivo metabolite-protein interactions revealed by large-scale systematic analyses. Cell 143(4): 639-650.
Article Information
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Category
Systems Biology > Proteomics > Whole organism
Microbiology > Microbial proteomics > Whole organism
Biochemistry > Protein > Interaction
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62 | https://bio-protocol.org/en/bpdetail?id=62&type=1 | # Bio-Protocol Content
Improve Research Reproducibility
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This is an In Press version of the protocol that has not yet been assigned to an issue.
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In-Solution Digestion Of Purified Yeast Protein For LC-MS
Xiyan Li
In Press
Published: Apr 20, 2011
DOI: 10.21769/BioProtoc.62 Views: 14782
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Abstract
This method describes the preparation of total yeast protein extract for mass spectrometry analysis. The protein extract is digested by trypsin in a solution with strong denaturants. The digested sample is dried and re-constituted in a mixture compatible with HPLC separation. Samples of isobaric labels should be processed in parallel experiments starting from trypsin digestion.
Keywords: Proteomics LC-MS Sample preparation In-solution digestion Yeast
Materials and Reagents
Purified protein
Sequencing grade modified trypsin (Promega Corporation, catalog number: V5113 )
6 M guanidine HCl
Tris HCl (pH 8.0)
1 M DTT
Triethylamine (TEA) (Sigma-Aldrich)
HPLC solvent A (usually 10% acetonitrile in water)
Acetic acid
Equipment
Amicon Ultra centrifuge filters Ultracel 10 k MWCO (EMD Millipore)
SpeedVac
Heat block
High Performance Liquid Chromatography (HPLC)
Amicon filters
Procedure
Concentrate purified protein on Amicon filters to 20 μl.
Take 20 μl protein solution (~100 μg), add to final of 6 M guanidine HCl, 50 mM Tris-HCl (pH 8.0), 2-4 mM DTT. Heat at 95 °C for 20 min.
Cool the reaction, then add 200 mM TEA. Final guanidine HCl concentration should be below 1 M.
Dissolve a vial of trypsin (20 μg) in 20 μl 50 mM acetic acid.
a. Add trypsin to target protein solution in a ratio of 1:50. Incubate at 37 °C for 1 h or longer.
b. SpeedVac the reation to dryness, then re-suspend with solvent A in HPLC.
References
Empirical lab protocol from Thermo Fisher.
Article Information
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© 2011 The Authors; exclusive licensee Bio-protocol LLC.
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Category
Systems Biology > Proteomics > Whole organism
Microbiology > Microbial proteomics > Whole organism
Biochemistry > Protein > Isolation and purification
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623 | https://bio-protocol.org/en/bpdetail?id=623&type=0 | # Bio-Protocol Content
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Metabolic Labeling of Yeast RNA with Radioactive Uracil
Yuehua Wei
Published: Vol 3, Iss 7, Apr 5, 2013
DOI: 10.21769/BioProtoc.623 Views: 9896
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Original Research Article:
The authors used this protocol in The EMBO Journal Aug 2009
Abstract
To examine gene expression, Northern blot or Real-Time PCR can be used to detect low abundant RNA such as mRNA. However, for high abundant RNAs such as rRNA and tRNA, Northern blot will not be able to discriminate the newly synthesized RNA from total RNA. Therefore, metabolic labeling is necessary to evaluate the expression of rRNA and tRNA genes. In this protocol, I describe a step-by-step method for labeling yeast RNA with radioactive uracil and examine the synthesis of these high abundant RNAs.
Keywords: Autoradiography Metabolic labeling Tritium RRNA TRNA
Materials and Reagents
Yeast strain of interest
RapidGel (500 ml) (Affymetrix, catalog number: 75848 )
Urea (CO(NH2)2) (Sigma-Aldrich, catalog number: U6504 )
Dextrose (C6H12O6) (Sigma-Aldrich, catalog number: G7021 )
Yeast nitrogen base without amino acids (Sigma-Aldrich, catalog number: Y0626 )
Synthetic dropout supplement without uracil (Sigma-Aldrich, catalog number: Y1501 )
Uracil (C4H4N2O2) (Sigma-Aldrich, catalog number: U1128 )
Rapamycin
TEMED/Tetramethylethylenediamine (C6H16N2) (Thermo Fisher Scientific, catalog number: 110-18-9 )
Ammonium persulfate (APS) ((NH4)2S2O8) (Sigma-Aldrich, catalog number: A3678 )
Formamide (CH3NO) (Thermo Fisher Scientific, catalog number: 75-12-7 )
DEPC/ Diethylpyrocarbonate (O(COOC2H5)2) (Sigma-Aldrich, catalog number: D5758 )
Bromophenol Blue (C19H10Br4O5S) (Sigma-Aldrich, catalog number: B0126 )
Xylene Cyanol FF (C25H27N2NaO6S2) (Sigma-Aldrich, catalog number: X4126 )
[5, 6-3H]-Uracil (PerkinElmer, catalog number: NET368250UC )
RNA marker (Promega Corporation, catalog number: G3191 )
Tris base (Thermo Fisher Scientific, catalog number: 77-86-1 )
Boric acid (Thermo Fisher Scientific, catalog number: 10043-35-3 )
EDTA (Sigma-Aldrich, catalog number: EDS-1KG )
Formamide loading dye
Small RNA separating gel
SD-Ura- (see Recipes)
SD-1/3 Uracil (see Recipes)
DEPC water (see Recipes)
10x TBE in DEPC water (see Recipes)
Small RNA separating gel (see Recipes)
2x Formamide loading dye (see Recipes)
Equipment
Bench top centrifuge
30 °C Shaker
Power supply with constant voltage > 450 V
Gel dryer (Bio-Rad Laboratories, catalog number: 165-1745 )
Exposure cosset/intensifier screen (Sigma-Aldrich, catalog number: C5479-1EA )
50 ml conical tubes
1.5 ml Eppendorf tube
Procedure
Inoculate yeast single colony in SD medium (or SD with appropriate dropouts). Shake 300 rpm at 30 °C overnight.
Dilute overnight culture in 50 ml conical tubes to 10 ml, OD600 =0.1 with SD-1/3 uracil and continue shaking until OD600 =0.4 (note: Reduction in cold uracil will allow hot uracil to be taken up by cells easily).
Cells were treated with drug and control vehicle, for example 100 nM rapamycin (final concentration) and its solvent methanol, continue shaking 300 rpm at 30 °C for desired time. In this specific experiment, rapamycin were added for 30 min.
Collect yeast cells by spinning down at 1,000 x g for 1 min at room temperature, remove supernatant (critical: Avoid putting yeast cells on ice. This is because ice will slow down growth, which will reduce significantly the uptake of hot uracil in the step 8 below).
Re-suspend yeast cells in 1 ml SD-Ura- (critical: Pre-warm medium to 30 °C), transfer to 1.5 ml eppendorf tube.
Spin down briefly by a bench top centrifuge at 5,000 x g, 15 sec, remove supernatant. Re-suspend yeast cells in 1 ml pre-warmed SD-Ura-.
Repeat 6 for 2 times and with the final re-suspension in 0.5 ml pre-warmed SD-Ura- (from the next step, collect radioactive liquid and solid waste in all steps, dispose according to environmental regulation).
Carefully add [5, 6-3H]-Uracil into each tube to the final concentration of 15 μCi/ml, vortex to mix, then put on a rack at 30 °C for 5 min.
Briefly spin down by a bench top centrifuge at 5,000 x g, 15 sec, remove supernatant.
Wash cells with SD-Ura-3 times as in step 5, ready to extract total RNA.
Total RNA was extracted by hot phenol method described in Wei (2012).
RNAs can be stored at -80 °C for up to 6 months.
Prepare ''small RNA separating gel'' on a large gel set (around 20 x 30 cm, mini gel did not work well).
Pre-run the gel for about 1 h at constant 450 V until the gel is heated to 50 °C.
Note: I found this step to be critical. One reason could be that pre-running the gel to this temperature could help get rid of excessive Urea in the gel, making RNA possible to go through. I usually attached a thermometer to ensure that the temperature has reached 50 °C.
Mix RNA samples with ''2x Formamide loading dye'' and heat at 70 °C for 2 min, then put samples on ice.
Turn off power. Rinse the wells with 1x TBE using a syringe with needle, make sure all urea is rinsed out from the wells.
Note: Urea is very dense and it will be impossible to load samples if urea is not rinsed out. If residual urea remains in the well, the resulting bands will be waving.
Load RNA samples (25 μg) with appropriate RNA marker and run gel at constant 450 V for about 2 h (BPB runs around 12 nt and cyanol around 55 nt).
Stain the gel with EtBr and take picture under UV light. This is total RNA which served as controls for newly synthesized RNA.
Sandwich the gel with autoclaved filter paper on one side and Saran-Wrap on the other side, put on a gel-dryer with filter paper side attach to the vacuum surface. Dry the gel at 80 °C for at least 2 h. Sometimes gels get cracked, may be because of insufficient drying or leaky vacuum.
The dried gel will stick to the filter paper. Wrap them in Sara-Wrap. 3H autoradiography in an exposure cassette with appropriate intensifying screen, for example Sigma Transcreen.
Develop after 4-7 days of exposure.
Recipes
SD-Ura-
Synthetic dextrose medium with Uracil dropout:
20 g Dextrose
1.7g Yeast Nitrogen Base
1.92 g synthetic dropout supplement without Uracil
5.0 g Ammonium Sulfate
Add ddH2O2 to 1 L and autoclave.
SD-1/3 Uracil: SD medium with 1/3 of regular Uracil
Similar to SD-Ura-except adding 25 mg Uracil.
DEPC water
Add 1 ml DEPC to 1 L ddH2O2, mix and put at room temperature overnight. Autoclave.
10x TBE in DEPC water
In 800 ml DEPC water, add:
108 g Tris base
55 g boric acid
40 ml of 0.5 M EDTA (pH 8.0)
Mix to dissolve and add DEPC water to 1 L. Autoclave.
Small RNA separating gel
Mix 2.5 ml 10x TBE, 6.25 ml RapidGel (40%) and 15 g Urea, heat to 50 °C and mix to dissolve.
Add DEPC water to 25 ml then filter through 0.45 μM Syringe.
Add 25 μl TEMED and 50 μl 25% APS, mix vigorously transfer to gel set with appropriate comb.
2x Formamide loading dye
95% (v/v) formamide in DEPC water, add tiny amount of Bromophenol Blue (0.01~0.1%) and Xylene Cyanol FF (0.01~0.1%), vortex to mix.
Acknowledgments
This protocol is derived from the following papers, Wei et al. (2009a) and Wei et al. (2009b), and the relevant references therein. The work was supported by NIH grants R01-CA099004 and R01-CA123391 to Dr. X.F. Steven Zheng at Robert Wood Johnson Medical School, University of Medicine and Dentistry of New Jersey.
References
Wei, Y., Tsang, C. K. and Zheng, X. F. (2009a). Mechanisms of regulation of RNA polymerase III-dependent transcription by TORC1. EMBO J 28 (15): 2220-2230.
Wei, Y. and Zheng, X. F. (2009b). Sch9 partially mediates TORC1 signaling to control ribosomal RNA synthesis. Cell Cycle 8(24): 4085-4090.
Wei, Y. (2012). A Simple Preparation of RNA from Yeast by Hot Phenol for Northern Blot. Bio-protocol 2(9): e209.
Article Information
Copyright
© 2013 The Authors; exclusive licensee Bio-protocol LLC.
How to cite
Readers should cite both the Bio-protocol article and the original research article where this protocol was used:
Wei, Y. (2013). Metabolic Labeling of Yeast RNA with Radioactive Uracil. Bio-protocol 3(7): e623. DOI: 10.21769/BioProtoc.623.
Wei, Y., Tsang, C. K. and Zheng, X. F. (2009a). Mechanisms of regulation of RNA polymerase III-dependent transcription by TORC1. EMBO J 28 (15): 2220-2230.
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625 | https://bio-protocol.org/en/bpdetail?id=625&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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Peer-reviewed
Northwestern Blot of Protein-RNA Interaction from Young Rice Panicles
Saminathan Thangasamy
Guang-Yuh Jauh
Published: Vol 3, Iss 7, Apr 5, 2013
DOI: 10.21769/BioProtoc.625 Views: 15480
Reviewed by: Tie Liu Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in The Plant Journal Jul 2012
Abstract
The northwestern assay is employed to study the interaction between protein and RNA. The RNA binding proteins tend to bind to different kinds of RNA through either known domains or unknown sequences of proteins. Rice LGD1 recombinant protein, a grass-specific novel protein with RNA binding sequences in its C-terminal, was used to probe its function as an RNA binding protein. The LGD1 comprising von Willebrand factor type-A domain (vWA), coiled-coil and nuclear localization signal (NLS) is a class of protein that localizes both in the nucleus and cytoplasm. Although LGD1 does not contain any putative RNA binding domains, we could find high-affinity RNA binding residues at the C-terminus using ‘RNABindR’ prediction software (Terribilini et al., 2007). The LGD1 recombinant protein, purified from bacteria, somehow forms both dimer and monomer even under denaturing conditions. However, only the dimeric form is able to bind to total and mRNAs. Due to its reproducibility and reliability, we believe that this protocol can be used across different organisms.
Keywords: LGD1 Multiple transcripts RNA binding Alternative splicing Rice
Materials and Reagents
Young panicle tissues of wild type (Oryza sativa L. ssp. Japonica cultivar Tainung67) plants
Tissues were collected from young panicles while they were in secondary branch initiation
Recombinant GST-LGD1 protein from E. coli (BL21)
pGEX-4T-1 GST expression vector (GE Healthcare, catalog number: 28-9545-49 )
Total RNA isolation kit - Trizol (Life Technologies, InvitrogenTM, catalog number: 15596-026 )
mRNA purification kit - Oligotex mRNA Mini Kit (QIAGEN, catalog number: 72022 )
RNA 5'-end Labeling kit (T4 Polynucleotide Kinase) (Thermo Fisher Scientific, catalog number: EK0031 )
FastAP Thermosensitive Alkaline Phosphatase (Thermo Fisher Scientific, catalog number: EF0651 )
[γ-32P]-ATP–for isotope labeling
PVDF Immobilon-P membrane (EMD Millipore, catalog number: IPVH15150 )
G-50 gel filtration dye terminator removal column (Geneaid, Taiwan, catalog number: CG050 )
Primary anti-GST antibody (Bioman, Taiwan, catalog number: GST001M ); HRP-conjugated secondary antibody (Thermo Fisher Scientific, catalog number: SA1-9510 )
SuperSignal West Pico Chemiluminescence detection system (Thermo Fisher Scientific, catalog number: 34087 )
Blocking buffer (see Recipes)
Equipment
Gel electrophoresis (Bio-Rad Laboratories, catalog number: 165-8005EDU )
SNAP i.d.® 2.0 protein detection system
Equipment for phosphor imaging (Bio-Rad Laboratories, catalog number: 170-9460 )
Procedure
RNA isolation and end-labeling of total and mRNA
Total RNA was isolated from young panicles of wild type plants with Trizol kit.
~0.5 g tissue was used to isolate ~500 μg of total RNA and 200 μg total RNA was used to purify ~2 μg mRNA using Oligotex mRNA Mini Kit. Nanodrop was used for final RNA quantitation (~2 μg/μl for total RNA; ~20 ng/μl for mRNA).
5 μg total RNA and 1 μg mRNA were first treated with FastAP Thermosensitive Alkaline Phosphatase (10 U) at 37 °C for 1 h to remove 5’-phosphate group according to manufacturer’s protocol.
The total and mRNAs were end-labeled with [γ-32P]-ATP using 1 μl (10 U) T4 Polynucleotide kinase at 37 °C for 30 min.
Labeled RNAs were separated from unincorporated label by gel filtration on Sephadex G-50 column.
Northwestern blot assay
GST-LGD1 fusion protein was extracted from E. coli harboring pGEX-4T-1 GST expression vector according to manufacturer’s protocol.
The proteins were separated on 12% SDS PAGE under denaturing conditions, and transferred to PVDF Immobilon-P membrane and detected using the SNAP i.d. protein detection system as follows.
For the Western blot, the primary anti-GST antibody (1:2,000) and secondary HRP (horseradish peroxidase)-conjugated antibody (1:3,000) were used for the enhanced chemiluminescence (ECL) detection.
SuperSignal West Pico Chemiluminescence was employed to detect western signals according to the manufacturer’s instructions.
For the Northwestern, recombinant proteins separated on 12% SDS PAGE under denaturing conditions were transferred to PVDF Immobilon-P membrane.
The proteins were then renatured on the blots overnight at 4 °C in a renaturation buffer containing 0.1 M Tris HCl (pH 7.5) and 0.1% (v/v) NP-40.
Blots were washed 4 times 15 min each in renaturation buffer and incubated 5 min at room temperature in blocking buffer.
Blots were hybridized overnight at 4 °C in blocking buffer (without BSA) in the presence of labeled total and mRNAs.
Finally, the blots were washed 4 times 5 min each in blocking buffer (without BSA and Triton) and autoradiographed to obtain the signal as seen in Figure 1.
Representative data
Figure 1. Northwestern blot analysis of LGD1 recombinant protein. The recombinant GST-LGD1 was purified from E. coli and analyzed through SDS-PAGE, western and northwestern blots. (a) Coomassie Brilliant Blue (CBB) staining shows two major bands (arrows). The size of the lower band is ~56 kD (monomer) and upper band is ~112 kD (homodimer). (b) Anti-GST antibody recognizes the GST-LGD1 fusion protein (arrows) as well as GST (open circle). Northwestern blot shows the total RNA. (c) and mRNA (d) isolated from young panicle tissues bind to upper band (arrows), but not to the lower band (open arrowhead). Asterisks indicate RNA binding to degraded form of LGD1.
Recipes
Blocking buffer
10 mM Tris HCl (pH 7.5)
5 mM Mg acetate
2 mM DTT
5 % (w/v) BSA
0.01 % (v/v) Triton X-100
Acknowledgments
We greatly appreciate the contributions of Drs Yue-Ie Hsing, Chyr-Guan Chern, Ming-Jen Fan and Su-May Yu to generate the TRIM database. We also thank Drs Ko Shimamoto (Nara Institute of Science and Technology, Japan), for sharing the pANDA-RNAi vector, and Su-May Yu (Institute of Molecular Biology, Academia Sinica), for excellent support with rice transgenic experiments. We also thank Ms Lin-Yun Kuang (Transgenic Plant Laboratory, Academia Sinica) for assistance in particle bombardment. We are grateful to Ms. Krisa Fredrickson for her English editing. This work was supported by research grants from Academia Sinica (Taiwan), the National Science and Technology Program for Agricultural Biotechnology (NSTP/AB, 098S0030055-AA, Taiwan), the National Science Council (98-2313-B-001-001-MY3, Taiwan) and the Li Foundation (USA) to Guang-Yuh, Jauh.
References
Uyttewaal, M., Mireau, H., Rurek, M., Hammani, K., Arnal, N., Quadrado, M. and Giege, P. (2008). PPR336 is associated with polysomes in plant mitochondria. J Mol Biol 375(3): 626-636.
Terribilini, M., Sander, J. D., Lee, J. H., Zaback, P., Jernigan, R. L., Honavar, V. and Dobbs, D. (2007). RNABindR: a server for analyzing and predicting RNA-binding sites in proteins. Nucleic Acids Res 35(Web Server issue): W578-584.
Thangasamy, S., Chen, P. W., Lai, M. H., Chen, J. and Jauh, G. Y. (2012). Rice LGD1 containing RNA binding activity affects growth and development through alternative promoters. Plant J 71(2): 288-302.
Uyttewaal, M., Mireau, H., Rurek, M., Hammani, K., Arnal, N., Quadrado, M. and Giege, P. (2008). PPR336 is associated with polysomes in plant mitochondria. J Mol Biol 375(3): 626-636.
Article Information
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© 2013 The Authors; exclusive licensee Bio-protocol LLC.
How to cite
Category
Plant Science > Plant biochemistry > Protein
Biochemistry > Protein > Interaction
Biochemistry > RNA > RNA-protein interaction
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